Evaluating strategies for integrating bacterial cells into a biosensor designed to detect electrophilic toxins
Katherine A. Linares
Thesis submitted to the Faculty of the
Virginia Polytechnic Institute and State University
in partial fulfilment of the requirements for the degree of
Master of Science
in
Environmental Engineering
Dr. Nancy G. Love, Chairperson
Dr. Brian J. Love
Dr. John C. Little
Dr. Kathleen Meehan
June 3, 2004
Blacksburg, Virginia
Keywords: biosensor, microbial stress response, potassium efflux, bacterial immobilization
Evaluating strategies for integrating bacterial cells into a biosensor designed to detect electrophilic toxins
Katherine A. Linares
Abstract
To improve the process stability of wastewater treatment plants, the construction
of a whole-cell bacterial biosensor is explored to harness the natural stress response of the
bacterial cells. The stress response selected in this work is the glutathione-gated
potassium efflux (GGKE) system, which responds to electrophilic stress by effluxing
potassium from the interior to the exterior of the cell. Thus, the bulk potassium in
solution can be monitored as an indicator of bacterial stress. By utilizing this stress
response in a biosensor, the efflux of potassium can be correlated to the stress response of
the immobilized culture, providing an early warning system for electrophilic shock. This
type of shock is a causative factor in many process upset events in wastewater treatment
plants, so the application of the sensor would be an early warning device for such plants.
The research conducted here focused on the biological element of the biosensor
under development. Three immobilization matrices were explored to determine the cell
viability and potassium efflux potential from immobilized cells: a calcium alginate, a
photopolymer, and a thermally reversible gel. The calcium alginate was unstable, and
dissolved after five days, such that the long-term impact of immobilization on the cells
could not be determined in the matrix. The photopolymer resulted in very low actvity
and viability of immobilized cellsOf the three matrices tested, indicating that the
composition of the polymer was toxic to the cells. Of the matrices tested, the thermally-
reversible gel showed the best response for further study, in that the matrix did not inhibit
cell activity or potassium efflux.
Acknowledgements
I would like to thank the funding sources that made this work possible: the Charles E.
Via, Jr. Department of Civil and Environmental Engineering Endowment, the Paul L.
Busch Award, and the EPA Midwest Hazardous Substances Research Center.
I would like to thank the members of my advisory committee, Dr. Nancy Love, Dr. Brian
Love, Dr. Kathleen Meehan, and Dr. John Little for their guidance and assistance
throughout my research.
I would like to thank our Laboratory Manager, Julie Petruska; Analytical Chemist, Jody
Smiley; and media man, Phil Wunderly for their assistance and guidance in the
laboratory.
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Table of Contents Introduction…………………………………………………………………………1 Chapter 1: Literature Review……………………………………………………….5 Introduction…………………………………………………………………5 Toxic shocks cause treatment upset………………………………………...5 Bacterial stress responses…………………………………………………...7 Real time information on wastewater toxicity………………………….….11 Immobilized cells as biological elements in biosensors…………………...12 Microfluidic sensors………………………………………………………..21 Matrices for immobilization………………………………………………..23 Scope of Research……………………………………………………….....26 References……………………………………………………………….....28 Chapter 2: Evaluating strategies for integrating bacterial cells into a biosensor designed to detect process upset Part A: viability and activity Abstract………………………………………………………………….…35 Introduction………………………………………………………………...35 Materials and Methods……………………………………………………..38 Results and Discussion……………………………………………....…….44 Conclusions………………………………………………………………...47 Acknowledgements……………………………………………………...…48 References………………………………………………………………….50 Chapter 3: Evaluating strategies for integrating bacterial cells into a biosensor designed to detect electrophilic toxins Part B: potassium efflux Abstract…………………………………………………………………….55 Introduction………………………………………………………………...55 Materials and Methods…………………………………………………….58 Results and Discussion…………………………………………………….64 Conclusions………………………………………………………………...67 Acknowledgements……………………………………………………...…67 References………………………………………………………………….69 Chapter 4: Engineering Significance………………………………………………80 Appendix A: Data for Chapter 2……………………………………………….....83 Appendix B: Data for Chapter 3…………………………………………………100
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List of Tables
Chapter 3: Evaluating strategies for integrating bacterial cells into a biosensor designed to detect electrophilic toxins Part B: potassium efflux Table 1. Efflux potential from immobilized cells……………………………66
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List of Figures Chapter 2: Evaluating strategies for integrating bacterial cells into a biosensor designed to detect process upset Part A: viability and activity Figure 1. Diagram of the OUR experiment protocol ………………………………52 Figure 2. OUR tests for cells immobilized in each polymer matrix….……………..53 Figure 3. Representative LIVE/DEAD® stains of immobilized cells ………………54 Chapter 3: Evaluating strategies for integrating bacterial cells into a biosensor designed to detect electrophilic toxins Part B: potassium efflux Figure 1. The glutathione-gated potassium efflux (GGKE) system………………...71 Figure 2. The experimental setup for potassium efflux experiments……………….72 Figure 3. Batch potassium efflux experiment from planktonic culture …………….73 Figure 4. Batch potassium efflux experiments across the growth curve for planktonic
cultures………………………………………………………….……….…...74 Figure 5. Effect of potassium on alginate stability…………………………..……...75 Figure 6. Potassium efflux from alginate-immobilized culture: Initial………..……76 Figure 7. Potassium efflux from alginate-immobilized culture: Day Five..……..….77 Figure 8. Potassium efflux from thermal gel-immobilized culture: Initial……….…78 Figure 9. Potassium efflux from thermal gel-immobilized culture: Day Three…….79
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Introduction
Process upset in wastewater treatment plants results in permit violations, low quality
effluent, and detrimental impacts on receiving waters. Even the most well-designed plant
may experiment occasional process upset due to shock loads of toxic and/or inhibitory
compounds. Unfortunately, the precise causes of these upset events are often unclear, and
due to the lack of real-time monitoring, operators are typically unaware that upset is
occurring until the biomass is already damaged. The implementation of a real-time sensing
system would provide operators with warning that a toxin is approaching and give them time
to proactively respond to reduce the impact of the shock, instead of merely reacting to an
upset event already underway.
In a survey performed by Love and Bott in 2000, operators reported that the most
common modes of process upset are ineffective biological oxygen demand (BOD) removal,
ineffective nitrification, deflocculation, and sludge bulking. Although the causes of these
upsets have been speculated, operators generally do not have sufficient information to
identify the source of the problem. However, when asked, operators suggested that heavy
metals and toxic organic compounds (which include electrophilic chemicals) are among the
most common sources of these upsets (Love and Bott, 2000). Thus, the major types of upset
events are thought to be due in part to electrophilic shock loads. Creating a sensor for
electrophilic compounds would provide warning of these types of upset events, although it
would not be useful for upsets due to other classes of compounds, such as oil and grease.
Because electrophiles elicit a natural stress response from bacterial cells, this stress
response can be harnessed to create a biosensor for electrophilic toxins. During an
electrophilic toxin upset event, the bacterial biomass initiates the glutathione-gated potassium
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efflux (GGKE) system (Bott and Love, 2002), in which potassium ions are effluxed from
within the cell (Apontoweil and Berends, 1975 a and b). By creating a biosensor with
bacterial cells containing the GGKE response, the intrinsic stress response of the bacterial
culture is harnessed and genetic engineering of the bacteria is not required. Many existing
biosensors have been slow to commercialize, mostly due to the limitations on genetically
engineered organisms. The commercial Microtox® assay, which uses naturally luminescent
Vibrio fischerii bacteria, is widely used as an indicator of toxicity, whereas many promising
biosensors with better response times and repeatability that use genetically modified bacteria
remain limited to laboratory use (for example, Philp et al., 2003; Gu and Choi, 2002).
Because the sensor developed in this work sensor does not require genetic modification, it
has an advantage in terms of eventual commercialization and field application.
By monitoring bulk potassium as an indicator of cellular stress, the biosensor will
monitor for electrophilic compounds that have been shown to cause upset in the activated
sludge process. The microscale biosensor under development includes a whole cell
biological element, potassium detection membranes, and microfluidic componentry that
compose a lab-on-a-chip device. The biological element consists of bacterial cells
immobilized in a hydrogel polymer matrix. Immobilization of the bacterial cells is important
because it keeps the cell concentration constant by maintaining the viability of the
immobilized cells while reducing cell division. Additionally, immobilized cells cannot
slough off as a biofilm layer could and clog the microfluidic channels in the device.
A series of screening experiments were conducted in this work to select an
appropriate polymer matrix for bacterial cell immobilization within the biosensor under
development, and are discussed in Chapter 2. The ideal matrix is physically and chemically
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stable over several weeks, non-toxic to the cells, and does not limit the diffusion of oxygen,
compounds that are necessary for cell growth, the electrophiles that would trigger a stress
response, or the potassium that would indicate that a stress response has been initiated.
Three matrices were examined here, including calcium alginate, several photopolymerizable
hydrogels, and a thermal polymer. The alginate showed good viability, but poor mechanical
stability, while the photopolymerizable polymers showed poor viability. The thermal
polymers were the best choice for continued study, in that the bacterial culture showed
extended viability after immobilization and the matrix did not inhibit the efflux response.
Experimental Objectives The overall objectives of this research were:
• Elucidate potassium efflux in response to a model electrophile from planktonic cultures of P. aeruginosa and E. coli at various growth states
• Select the growth state and the strain with maximal potassium efflux per cell for
further study
• Screen a series of immobilization hydrogels for cell viability within the matrix
• Quantify potassium efflux from immobilized bacterial culture selected The specific results and methods used to meet these objectives will be discussed within the
following chapters of the thesis.
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References Apontoweil, P., Berends, W. (1975a). Glutathione biosythesis in Escherichia coli K-12: properties of the enzymes and regulation. Biochimica et Biophysica Acta, 399 (1), 1-9. Apontoweil, P., Berends, W. (1975b). Isolation and initial characterization of glutathione-deficient mutants of Escherichia coli K-12. Biochimica et Biophysica Acta. 399(1), 10-22. Bott, C. B. and Love, N. G. (2002) Investigating a mechanistic cause for activated sludge deflocculation in response to shock loads of toxic electrophilic chemicals. Water Environment Research. 74:306-315. Gu, Man Bock, and Sue Hyung Choi. A portable toxicity biosensor using freeze-dried recombinant bioluminescent bacteria. Biosensor and Bioelectronics. 17 (2002) 433-440 Love, N. G., Bott, C.B. (2000). WERF Project 99-WWF-2 Report: A review of and needs survey of upset early warning devices. Water Environment Research Foundation. Philp, Jim C., Severine Balmand, Eva Hajto, Mark J. Bailey, Siouxsi Wiles, Andrew S. Whiteley, Andrew K. Lilley, Janos Hajto, and Sandra A. Dunbar. (2003). Whole cell immobilized biosensor for toxicity assessment of a wastewater treatment plant treating phenolics-containing waste. Analytica Chimica Acta.
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Chapter 1: Literature Review
Introduction
Process upset events in wastewater treatment plants result in high nutrient and
pathogen loading to receiving waters, endangering public heath and stability of waterways.
Electrophilic compounds are one cause of upset events, typically resulting in deflocculation,
which is thought to be due to potassium efflux through the glutathione-gated potassium
efflux (GGKE) mechanism (Bott and Love, In Press). Controls to eliminate such upset
events are limited by the scarcity of real-time toxicity monitoring devices. Biosensors show
promise in the field and have been developed to respond to specific compounds (for example,
Strachan et al., 2001 and Reid et al., 1998) as well as general stress and oxidative damage
(for example, Belkin et al., 1997 and Elasri et al., 2000). Most biosensors employ
genetically modified bacteria, which limits their potential for commercialization. Non-
engineered strains have been used to some degree, but more work is needed in this area. To
create a real-time live cell sensor, a flow-through system is required. Microfluidic
technology can be applied to create micro-scale sensors containing bacteria immobilized in
hydrogels (for example, Heo et al., 2003), because these sensors provide rapid response and
require low sample volumes. Incorporating these microfluidic sensors into a real-time
monitoring system can provide forewarning of toxic shock loads to treatment plants,
allowing operators time to proactively mitigate the threat.
Toxic shocks cause treatment upset
Wastewater treatment plants employing biological treatment schemes to reduce or
remove organic and inorganic pollutants play a pivotal role in pollution reduction. The
5
optimal functioning of these plants is critical for receiving stream health and downstream
water reuse capacity. Despite the dependence on these plants for pollutant removal, even the
most well designed plant may experience occasional process upset events due to changes in
flow pattern or waste composition, or due to toxic loads of inhibitory chemicals.
A survey of Water Environment Research Foundation subscribers revealed that the
common modes of process upset are ineffective BOD removal, deflocculation, ineffective
nitrification, and sludge bulking (Love and Bott, 2000). Some survey responders reported
that the most common causes of upsets they experienced were high flow, toxic organics, oil
and grease, and heavy metals (Love and Bott, 2000). Unfortunately, in most cases, the cause
of the upset is commonly not discovered until well after the event, if at all. Interestingly,
several of the purported causative factors can be grouped as electrophiles, such as the heavy
metals and selected toxic organics.
During process upset from a shock load of an electrophilic compound, the treatment
plant loses treatment efficiency because the chemical stressors cause deflocculation of the
activated sludge flocs. This loss of biomass from the system is believed to occur because
stress response mechanisms cause bacteria to efflux potassium (Bott and Love, 2002). The
additional monovalent cations in the floc matrix is believed to increase the intrafloc divalent
to monovalent cation ratio (Bott and Love, 2002), which is an important predictor of floc
strength (Higgins and Novak, 1997a and 1997b). Disrupting this balance causes the flocs to
lose stability and results in loss of biomass from the system (Murthy et al., 1998).
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Bacterial stress responses
Oxidative stress mechanisms in bacteria
Damaging oxidative (electrophilic) chemicals enter cells either from the outer
environment or are generated internally as part of cellular metabolism. The types of
oxidative stressors generated within the cell (reviewed in Storz and Imlay, 1999) are
superoxide anions, hydrogen peroxide, and hydroxyl radicals, as well as methylglyoxal
(Kalapos, 1999). Superoxide anions, O2•-, are reduced to O2 and H2O2 by superoxide
dismutase. Hydrogen peroxide originates from the reduction of superoxide by its dismutase
or by reduction of oxygen. Hydroxyl radicals (HO•) are produced from superoxide and
H2O2, and are very reactive with and damaging to DNA. Example equations illustrating
these reactions (reviewed in Storz and Imlay, 1999) are provided below. Equation 1 shows
the complete stepwise reduction of the superoxide radial through peroxide and hydroxyl
radical to water. Equation 2 shows the reduction of the oxygen radial to peroxide, a reaction
catalyzed by superoxide dismutase. Equation 3 shows the reduction of peroxide to water, a
reaction catalyzed by catalase.
(1) O2 O2•- H2O2 HO• + H2O H2O
(2) O2•- + 2 H2 H2O2 + O2
(3) 2 H2O2 O2 + 2 H2O
The types of damage caused by oxidative stress include DNA damage and mutagenesis, as
well as membrane and protein damage.
The presence of these electrophiles changes the intracellular oxidation state. Two
genes, oxyR and soxR, exist to detect such shifts in redox potential. Redox regulation is
defined as the control of protein activity by oxidation and reduction and is a major means of
7
cellular control (as reviewed in Demple, 1996 and Pomposiello and Demple, 2001). The first
step in protecting cells against damage is sensing the oxidants, achieved by the proteins
OxyR and SoxR. These proteins are bound to the promoter region of the genes under their
control such that the conformational change occurring upon oxidation allows transcription of
these response genes (as reviewed in Pomposiello and Demple, 2001).
As reviewed in, for example, Storz and Imlay (1999) and Pomposiello and Demple
(2001), OxyR is activated and deactivated by the formation and reduction of disulfide bonds.
Peroxide exercises direct control over the activation of OxyR by reducing pairs of sulfur-
hydrogen bonds to disulfide bonds. When oxidized by an electrophilic compound, the OxyR
protein self-regulates the oxyR gene, as well as dps (protein to bind iron and DNA), gorA
(GSH reductase), grxA (glutaredoxin), katG (peroxidase), ahpCF (NADPH reductase), and
fur (iron binding protein to prevent iron transport). Additionally, through its control of oxyS
and production of OxyS protein, OxyR controls other regulatory genes such as rpoS and flhA.
The majority of the information on the OxyR system has been obtained from E. coli, but
homologs have been found in many other species. After oxidation, OxyR is returned to the
reduced state by the GSH-glutaredoxin-1 (grxA) system (as reviewed in Storz and Imlay,
1999 and Pomposiello and Demple, 2001).
The SoxR protein is constitutively expressed at a minimal level within the cell and
functions through the activity of its [2Fe-2S] centers, which can be oxidized and reduced by a
one-electron shift (Ding and Demple, 1997) as reviewed in, for example, Storz and Imlay
(1999) and Pomposiello and Demple (2001). Superoxide exercises direct control over the
activation of SoxR by shifting the oxidation state of Fe+ to Fe2+ in the iron-sulfur centers.
The controlling compound for oxidation may be superoxide or depletion of NADPH. The
8
SoxR responds in the presence of heavy metals, antibiotics, and organic compounds, all
superoxide-generating compounds, and also responds to nitric oxide (Zheng and Storz,
2000). When activated by an oxidant, SoxR initiates expression of the soxS gene. The genes
under control of the soxRS regulon are sodA (superoxide dismutase to deactivate oxidants),
zwf (glucose-6-phosphate dehydrogenase to recreate the NADPH pool), fldA and fldB
(flavodoxins to reduce metals in constituent groups on intracellular molecules), fpr (NADPH
reductase), fur (iron binding protein to prevent iron transport), nfo (DNA repair gene), arcAB
(efflux pump to remove toxic chemicals), and micF (reduces porin expression and thus cell
permeability). All of these genes that are controlled by SoxR work to prevent or mitigate
damage caused by oxidants. These genes were elucidated in E. coli, but many other bacterial
strains contain SoxR homologs (as reviewed in Storz and Imlay, 1999 and Pomposiello and
Demple, 2001).
Another type of intracellular oxidative stress arises from methylglyoxal, an
electrophilic byproduct of metabolism, also called pyruvaldehyde, pyruvic aldehyde, 2-
oxopropanal, 2-ketoproprion-aldehyde, or acetyl-formaldehyde. Methylglyoxal generation
occurs during upswings in the electron donor concentration, accumulating in starving cells
that are suddenly exposed to high concentrations of carbon source. The pathways for glucose
oxidation become overwhelmed, and methylglyoxal is formed to release some of the excess
carbonaceous energy (Kalapos, 1999). It has been shown (Ness et al., 1997) that the
glutathione-gated potassium efflux system evolved as a mechanism of protecting the cell
from electrophilic damage from methylglyoxal.
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Glutathione-gated potassium efflux (GGKE)
Reduced glutathione (GSH) (N-(N-L-γ-glutamy-L-cysteinyl)glycine) is constitutively
expressed in the cell (reviewed in Storz and Imlay, 1999). Due to the activity of its
sulfhydryl group, GSH acts in a sacrificial role to bind electrophiles and oxidizes into GSSG,
as well as oxidized glutathione-S-conjugates (GSX) (Apontoweil and Berends, 1975 a and b).
GSX triggers the cell stress response of potassium efflux through the glutathione-gated
potassium efflux (GGKE) mechanism controlled by the KefB and KefC membrane transport
proteins (Apontoweil and Berends, 1975 a and b; Booth et al., 1993; Elmore et al., 1990,
Ferguson et al., 1995; Munro et al., 1991). GGKE shuttles potassium ions out of the cell in
an ion exchange for hydrogen into the cell. The transport of these hydrogen ions dissipates
the proton gradient, but acidifies the cytoplasm, causing the DNA to supercoil protecting it
from oxidative damage (Ferguson et al., 2000).
The glutathione control system is involved in the OxyR stress response, in that OxyR
is reduced by enzymatic reaction with glutaredoxin 1 (Grx1), produced through OxyR
control to reduce oxidized glutathione back to the reduced form, GSH (Zheng et al., 1998).
Because the production of Grx1 is regulated by OxyR, the reduction of OxyR is
autoregulated. Zheng and Storz (1998) presented a conceptual equation for this reaction
obtained through both molecular and biochemical studies:
OxyR (ox) + 8 GSH OxyR (red) + 4 GSSG
Interestingly, OxyR is reduced by sacrificially oxidizing GSH into GSSG (Zheng and Storz
1998). Further, the redox potential of OxyR was found to be –185 mV, much higher than the
typical unstressed cell redox potential, indicating that OxyR is primarily in the reduced form
normally (Zheng and Storz, 1998).
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Overall, bacteria respond to electrophilic stress, detected by OxyR, SoxR, and
glutathione, by a number of protective mechanisms. The GGKE system is linked to the
OxyR system through the reduction of the oxidized proteins back to their reduced forms.
Although SoxR and OxyR control many intracellular responses, such as the activation of
protective genes, the GGKE system differs in its interaction with the Kef potassium efflux
channels because the system triggers an observable extracellular effect. The efflux of
potassium appears to cause deflocculation in wastewater sludge flocs, a typical symptom of
process upset caused by toxic shock.
Real time information on wastewater toxicity
The key to successful control of toxic shock events is awareness of their impending
arrival at the plant as well as the causative toxin and its potential effect. Recognizing that a
threat is approaching gives the operators time to respond in order to mitigate the threat.
Control of wastewater plants is typically performed by operators with the help of
computerized systems such as the Supervisory Control and Data Acquisition (SCADA)
system. However, these systems are limited by the lack of real-time monitoring devices (for
example, Dieu, 2001). Parameters that can be evaluated quickly include flowrate, pH,
temperature, and dissolved oxygen, which are inadequate to accurately predict plant
functionality (Bungay and Andrews, 1970, Olsson and Andrews, 1981). The five-day BOD
test clearly takes too much time to be helpful in identifying problems in treatment. Even the
chemical oxygen demand (COD) test, which can be completed in a few hours, is too long to
wait to mediate an upset because the upset has already begun by the time the effluent COD
spike is seen. Therefore, these tests fail as process control techniques. One simple way to
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use existing technology to improve process control is the use of the dissolved oxygen (DO)
profile. The variations in DO along the length of the biological reactor comprise the DO
profile. Monitoring this profile permits operators to reduce the energy used to aerate the tank
and to estimate the oxygen uptake rate. Shifts in the uptake rate as an alteration in the
respirometry response can signal process upset (Olsson and Andrews, 1981, Bergeron and
Paice, 2001).
Most control systems use “feed-back” setups, in that data from the effluent
determines if treatment is effective (Bungay and Andrews, 1970). In order to effectively
control plant function, “feed-forward” control is necessary to alert operators to potential
problems, which would link real-time on-line sensors to global control systems within the
plant (Bungay and Andrews, 1970). The limitation on these control systems is the lack of
real-time sensors for causative factors in process upset. Lab analyses of the components in
the influent are time consuming and costly, particularly when the target compound is
unknown. The use of biosensors to provide real-time feedback on toxicity or process
conditions shows promise for improving process control in wastewater treatment.
Immobilized cells as biological elements in biosensors
A biosensor is generally defined as an analytical tool that pairs a biological
component with a detection system to produce a measurable response, which can be
quantified into the amount of analyte present in a sample (Belkin, 2003). The response is
often measured electrochemicalyl or optically. The types of biological components that can
be used are enzymes, antibodies, proteins, nucleic acids, or whole cells. Whole cells are less
expensive to use because their components require no isolation and purification, as in the cell
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component assay systems. Also, whole cells provide information on the bioavailability of
the toxins within the sample instead of simply reporting the concentration present. However,
whole cells have a certain limited lifespan, require longer response times than isolated
components, and may show significant variability between cell batches (Ramanathan, 2000).
Reporter genes
Much work to date with whole cell biosensors has focused on genetically engineering
bacterial strains to elicit specific responses to particular compounds. Three types of reporter
genes are typically used: lux, luc, and gfp. First, bacterial luciferase (lux) produces the
oxyluciferin protein by combining the substrate luciferin with oxygen in the presence of the
luciferase enzyme. The oxyluciferin product is in the excited state and produces a photon,
which can be detected and used to quantify the response (Hastings, 1983). The yield of this
reaction is low, only about 0.1 yield units, and it consumes reducing power, the precursor to
ATP formation. Further, the reaction requires a substantial amount of oxygen to proceed,
limiting its use to aerobic conditions only (Lewis et al., 1998). The lux gene was originally
isolated from three bacterial species: Vibrio and Photobacterium, both marine, and
Photoorhabdus, a terrestrial species (Keane et al., 2002). Second, firefly luciferase (luc) was
isolated from the beetle Photinus pyralis and also produces oxyluciferin, but requires a two-
step enzyme reaction and consumes ATP. This reaction has a yield of 0.88 (Lewis et al.,
1998) and a wide linear range. However, the reaction only proceeds below 30oC (Keane et
al., 2002), so it is less frequently used in bacterial work, as the cells typically require higher
temperatures for optimal growth. Third, green fluorescent protein (gfp) from the jellyfish
Aequorea victoria requires a much smaller amount of oxygen than luc or lux and the light
13
chromophore is directly incorporated within the crystal structure. Production of the response
protein is much slower than the luciferase-based systems (Keane et al., 2002).
The selection of the reporter gene for a particular application is typically based on the
desired time response of the reporter. Luciferase systems, in which the fluorescent proteins
are broken down in a short time, are used for time-based assays in which repeated
measurements are needed. Green fluorescent protein systems are best for single
measurement studies where one yes or no answer is desired because hours are required to
produce and properly fold the protein. Additionally, once generated, the fluorescent signal
remains robust for much longer than the luciferase signals, resulting in a longer measuring
window with a constant signal output (Keane et al., 2002).
Two methods exist for adding response genes to cells, which can be classified as
“lights on” and “lights off”, or induced and constitutive. Inserting the reporter gene into a
general cell maintenance section of the genome will result in constant production of green
fluorescence, while reduced intensity indicates some type of insult has occurred (Belkin,
2003). Alternately, the reporter can be linked with a promoter that regulates gene expression
of a desired reporting characteristic, such as DNA repair. In this setup, an increase in light
level functions as the indicator (Belkin, 2003).
Genetically engineered strains in biosensors
Several groups have used genetically modified strains to create biosensors specific to
certain chemicals. A biosensor using Pseudomonas aeruginosa containing the lux operon to
respond to polycyclic aromatic hydrocarbons (PAHs) was created by immobilizing the cells
in poly(vinyl alcohol) (Philp et al., 2003), which showed good repeatability on standards but
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variable results on real samples. Also, E. coli, P. fluorescens, and P. putida with a lux
reporter gene were used to construct a biosensor to respond to PAHs via bioluminescence
(Reid et al., 1998) and, although real samples were not tested, the sensor showed a stable,
linear, and dose-dependent response to standards. A heavy metal responsive plasmid was
constructed using luc to respond with production of luciferase and inserted into
Stapholococcus aureus and Bacillis subtilis (Taurianinen et al., 1998). This sensor
responded within three hours to lab samples and responsitivity was not reduced by freeze-
drying the cells before use. Kohler et al. (2000) used E. coli modified with the lux operon to
make a biosensor for detecting 4-chlorobenzoic acid by immobilizing cells in alginate in a
microtiter plate, but more work was needed to optimize the system before real samples can
be tested. Strains of Pseudomonas and Archromobacter were used to perform preliminary
work towards creating a biosensor for surfactants using amperometric detection (Taranova et
al. 2002). Gu and Choi (2002) used freeze-dried E. coli with a reporter bound to the phenol
degradation pathway to create a biosensor for phenol detection in a complete portable field-
ready kit. A range of phenols were tested with rapid, repeatable results, but no results from
field samples were reported.
Hansen et al. (2000) used E. coli to create a biosensor for detecting mercury by fusing
the Pmer, a promoter induced by Hg, and the mer reporter gene for Hg to luxCDABE, lacZYA,
and gfp. These constructs produce luminescence, beta-galactosidase, and green fluorescent
protein, respectively. Each construct was cloned into a delivery vector to permit its insertion
into any bacterial strain desired. P. aeruginosa was used as an example. Other groups
(Ramanathan et al., 1997; Lyngberg et al., 1999; Rasmussen et al., 2000; Selifonova et al.,
1993) had performed similar research with the mer reporter gene encoded on a plasmid, but
15
mixed results were obtained because the plasmids were unstable. However, Petanen and
Romantschuk (2002) used plasmids with a luc response to mercury and arsenite in P.
fluorescens and obtained results comparable to traditional analytical methods, but much
faster.
Howbrook et al. (2001) created a whole cell biosensor for glucose using E. coli with a
luminescence reporter bound to the katG gene, which is tightly controlled by the OxyR
system such that it responds to oxidative stress. As proof of concept, the glucose oxidase
enzyme was added to the sensor to produce H2O2 when glucose is present, thus creating
oxidative stress within the cell to trigger the OxyR response system and initiating the lux
reporter. Any such oxidase enzyme could be included to target sensor specificity for a wide
variety of compounds. The assays were conducted in liquid culture with samples generated
in the laboratory.
Creating a general response biosensor is typically approached by developing a panel
of bacteria, each responsive to a different threat. Panels of E. coli bacteria were engineered
to create sensors for heat shock, oxidative stress, and protein damage (Belkin et al., 1997),
DNA damage (Belkin, 2003), oxidative damage, membrane damage, DNA damage, and
protein damage (Kim and Gu, 2003), and heat shock, oxidative stress, fatty acids, peroxides,
and genotoxicity (Premkumar et al., 2003). A lux-containing E. coli strain immobilized in
poly (vinyl alcohol) (PVA) was used to create a slow release biosensor for general stress
where the PVA matrix was dissolved using KCl over time so that new cells were being used
for each toxicity test (Horsburgh et al., 2002). Biosensors for genotoxicants were created by
inserting a lux reporter into the recA promoter region for DNA damage repair in E. coli.
16
(Polyak et al., 1997; Belkin, 2003; Polyak, 2001). Elasri et al. (2000) created a similar
sensor using P. aeruginosa.
Two well-known assays for genotoxicants and cytotoxicants are the SOS-Lux test and
the Lac-Fluoro test. The SOS-Lux test was created by adding the lux reporter under the
control of the SOS DNA damage repair system to identify genotoxicants. The Lac-Fluoro
test uses constitutive expression of gfp, which is inserted to report cell maintenance function
and to indicate cytotoxins. Decreases in fluorescence shows cytotoxicity, or impacts to cell
metabolism and function (Baumstark-Khan et al., 2001).
The toxicity of heavy metals (chromium, zinc, copper, nikel, and arsenic) was
evaluated using a microtiter plate setup and two strains of Salmonella typhimurion, one
modified for the SOS-Lux and one for the Lac-Fluoro test (Rabbow et al., 2002).
Improvements to the sensitivity of the SOS-Lux test were obtained by inserting the reporter
plasmid into E. coli containing a tolC mutation, which made the membrane more permeable
to hydrophobic substances. The sensitivity was further improved by using a S. typhimurion
strain with a defect in the cell membrane, which again increases the membrane permeability
(Rettberg et al., 2001). The SOS-Lux and the Lac-Fluoro tests developed by Rabbow et al.
(2002) have been applied in practice on the international space station to determine toxicity
of recycled water supplies on the station. A different sensor formulation with the similar
genetic modifications has been used to determine the threat from radiation on the space
station (Rabbow et al., 2003).
On the whole, genetically engineered strains used in biosensors have not been
commercialized to a large degree. The difficulty in overcoming the genetically modified
organism (GMO) regulations makes such sensors primarily useful only in laboratory settings,
17
where the threat of accidental release is not applicable. Despite their advantages in terms of
robust, repeatable responses, engineered strains have yet to be widely commercialized.
Non-engineered strains
One drawback of the majority of work to date is that most sensors employ genetically
modified bacteria as the reporter. A notable exception is the commercial Microtox® assay,
which uses naturally luminescent Vibrio fischerii bacteria. Non-engineered cells do not
mandate such strict controls on release as those that have genetic modifications. While
engineered cells may yield a more robust response, wild-type cells facilitate eventual
commercialization of a sensor. Additionally, the use of indigenous wild type cells more
accurately reflects the conditions under observation than the use of an engineered lab strain.
Using wild-type cells means that the intrinsic cell processes must be harnessed to detect the
compounds of interest or to determine if the cells are inhibited.
BOD sensors typically do not require genetic engineering to create a functional sensor
since the concentration of oxygen in a sample can be easily monitored. These sensors are
typically constructed using an immobilized bacterial culture contained between a dialysis
membrane and a gas permeable membrane. The sample fluid diffuses through the dialysis
membrane and passes through the culture where some of the oxygen in the sample is
consumed. The amount of oxygen remaining in the sample is measured using a dissolved
oxygen electrode placed above the gas permeable membrane. The respiration rate of the
culture is calculated from the difference in oxygen concentrations between the amount in the
sample initially and that detected by the electrode after exposure to the cells (Liu and
Mattiasson 2002). Current work (Heim et al. 1999) has focused on combining two pure
18
cultures to expand the limited range of compounds detected when one pure culture is used, or
an ensemble of bacteria, which shows unstable response over time as different species
dominate (Tan and Wu 1999). In general, these sensors are complicated to apply because the
respiration rate shifts depending on the amount of substrate present in the sample. Adjusting
for this variation, this type of BOD sensor has been shown to be as effective as the traditional
five-day BOD test but with a response within minutes. However, the variability between
sensors of the same construction is significant (Liu and Mattiasson 2002).
In addition to BOD sensors, amperometric biosensors have been developed that
function by binding cells to a screen printed electrode to measure the transfer of electrons
during substrate consumption. One example of this type of sensor was developed by Skladal
et al (2002) to create a phenol sensitive biosensor. Pseudomonas strains were immobilized
on a screen-printed electrode such that the uptake of phenol as a carbon and energy source by
the cells can be detected by the electrode as electrons are transferred. The sensor showed a
rapid, reproducible response. However, the sensing surface had to be prepared and used on
the same day and the response varied depending on the ionic strength of the sample.
Natural products of cells can also be monitored to determine if they are inhibited. For
example, Guven et al. (2003) created a bioassay for inhibition of enzyme biosynthesis to
monitor the concentrations of two enzymes, one produced by E. coli and the other produced
by Bacillus subtilis. Colorimetric tests were used to monitor beta-galactosidase synthesis in
E coli, which is created during consumption of glucose and is metabolism related, and alpha-
amylase synthesis in B. subtilis, which is created during late log stage and is growth phase
related. Changes in the concentrations of these enzymes indicated the presence of toxic
compounds. The system worked very well for pesticides, the model organics tested, but was
19
less repeatable with heavy metals. This type of assay takes much longer than an engineered
“lights on” or “lights off” strain, because up to eight hours are required to determine if a
decrease has occurred in the enzyme synthesis.
Cell components in biosensors
The use of cell components (purified enzymes) in biosensors is an attractive
alternative to whole, living cells, which require a constant supply of nutrients and oxygen.
These systems typically consist of the sensing element (enzyme) bound to an electrode such
that the change in concentration of a product is converted into an electric signal monitored by
a detector (reviewed in Karube and Nomura, 2000). An example of such a system was
created by Moser et al. (2002) in which oxidase enzymes specific to glucose, lactate,
glutamate, and glutamine were immobilized into a flow through microfluidic device. The
response was rapid and repeatable and a single sensor could perform continuous monitoring
of all four compounds of interest. However, isolation of the enzymes used in this type of
sensor is costly and time consuming and enzymes alone are less stable than whole cells
(Karube et al., 1995).
Using whole cells that have been killed provides a useful method of preserving the
enzymes and eliminates the purification step. Tan and Zhenrong (1998) used thermally
killed Bacillus subtilis cells immobilized in a membrane bound to the end of a DO probe to
create a sensor for BOD. The BOD reported by the sensor correlated well with BOD5 assays
performed on identical samples, with results available within half an hour. This type of
biosensor functions by incorporating the active components necessary to catalyze the
20
oxidation of carbon sources, specifically, enzymes and cofactors, but without including the
difficult and time consuming step of isolating these enzymes from living cells.
Microfluidic sensors
These macroscale sensors using whole cells or cell components can be improved by
miniaturization. Reducing the size of a sensor to the microscale to create
microelectromechanical systems (MEMS) presents several advantages in sensing ability.
Microsensors require much lower sample and analyte volumes, present a more rapid response
than full-scale sensors, and can be made disposable because of the small size and volumes
involved, thus creating a highly cost-effective analysis system. However, the robustness of
these systems to harsh environments has not been proven and they have also been slow to
commercialize.
Several groups have used microfluidic technology to create biosensors. One
prominent example relavent to environmental engineering is the class of BOD sensors
created by Yang et al. (1996, 1997), where whole cells were immobilized directly onto the
surface of a microscale DO probe. The cells used were Trichosporon cutaneum, useful for
BOD sensors because they consume a wide range of carbon sources, not a limited few
compounds. In this way, they report a BOD closer to the BOD5 than what would be reported
by a sensor using a strain such as Pseudomonas, which prefers to grow on acetate. With the
sensor, this group was able to achieve sensor BOD to BOD5 ratios of 0.65 to 1.70 with real
wastewater samples and a repeatability of plus or minus eight percent was reported. The
response time was twenty minutes and the sensor was linear for a BOD range of 0.2 to 18
mg/L BOD. Additionally, because it is on the microscale, the device is smaller and easy to
21
handle. Such a sensor would be useful to indicate spikes in influent and effluent BOD; for
such an application, the limited precision of the device is less of a factor because the purpose
of the sensor is to provide nearly real-time monitoring, not reportable BOD values.
In addition to whole cells, enzymes are commonly used within the microscale sensors
as reporters for the presence of certain compounds. Zhan et al. (2002) immobilized a pH
reporting dye and enzymes to catalyze specific reactions in a microreactor within a
microfluidic channel array to create a biosensor for potentially any compound of interest.
Adding the correct enzymes created a pH shift, reported by the dye, and the magnitude of the
shift correlated to the concentration of the analyte that reacted with the enzymes. Moser et
al. (2002) created a bio-MEMS using enzymes bound within a flow channel with electrodes
to report changes in current as the enzymes reacted with the species of interest, which were
glucose, lactate, glutamate, and glutamine. Petrou et al. (2002, 2003) created a bio-MEMS
sensor for glucose and lactate by immobilizing enzymes in a microchannel and measuring the
response with a dialysis probe made of polyacrilonitrile fiber. Zimmerman et al. (2003)
created a flow-through glucose sensor by immobilizing glucose oxidase enzyme in polyvinyl
alcohol (PVA) polymerized by UV light within a microfluidic system. Due to high
fabrication temperatures associated with the anodic bonding between wafer sheets, it was
important that the enzymes were immobilized in situ after device fabrication.
These types of sensors were designed for medical applications, such as monitoring
the concentrations of sugars in blood samples. Although the concept could be modified for
application in environmental engineering processes by changing the enzymes used in the
sensors, this design is less useful for general monitoring. Such sensors are primarily useful
for monitoring the concentration of a few known compounds, and so could be applied to
22
sensing compounds known to be present in environmental samples. However, such sensors
do not address the need for monitoring a wide range of toxic compounds.
Immobilization Matrices
The immobilization of cells within a microfluidic sensor is critical to the functionality
of the system. Using a biofilm of cells adhered to the bottom of the flow channel is likely to
be unstable, as cells and chunks of film can slough off with time and foul the detector or clog
the channel. Immobilizing cells within hydrogels prevents sloughing, provides some degree
of protection for the cells, and exposes the cells to an environment similar to that within a
biofilm (Junter et al., 2002). There are two primary types of hydrogels, those that are
chemically polymerizable and those that are photopolymerizable.
Chemically polymerized hydrogels, such as sodium alginate, are useful for
macroscale work by forming beads from the gel precursor solution (Drury et al., 2004).
However, downsizing the bead to microscale may prove difficult to form. Therefore,
photopolymerized polymers are typically used for microfluidic applications because they can
easily be formed into a variety of shapes in situ by using various photomasks to control what
areas of the polymer precursor are polymerized. First, Beebe et al. (2000) created various
shapes of hydrogels by polymerizing acrylic acid and methacrylate in situ in a microchannel,
showing the ease of handling with the photopolymer. Further, this group used pH-sensitive
polymers to control flow within a microfluidic channel matrix by placing polymers of
varying compositions at critical points in the flow matrix such that shifts in pH would cause
expansion or contraction of the polymer patch and, thus, either allow or block flow through
particular regions of the channels. This passive method of flow control simplifies
23
microfluidic componentry, but for biological systems, the pH extremes required to change
the flow regime would prove detrimental to the cells in the system. Moving from flow
control to the development of a reporting system, Zhan et al. (2002) immobilized the glucose
oxidase enzyme within several micropatches in a microfluidic device. The device reported
the concentration of glucose by shifts in the intensity of the reporter dye co-immobilized with
the enzyme. This sensor would be used for medical applications, but still shows the
usefulness of the photopolymer in microfluidic work.
Enzymes as well as viable cells have been immobilized within photopolymers. Koh
et al. (2002) encapsulated mammalian cells within polyethylene glycol (PEG) diacrylate
(PEG-DA) with Darocure 1173 as the photoinitiator polymerized with UV light. Cells were
viable up to a week after polymerization based on observations using the LIVE/DEAD
viability/cytotoxicity fluorescence staining. This application was for tissue engineering and
not biosensor development. However, Heo et al. (2003) immobilized E. coli cells in PEG-
DA within a microfluidic channel constructed of polydimethylsiloxane (PDMS) using
photopolymerized micropatches of gel spanning the height of the channel, but not the width,
to permit flow around the edges of the hydrogels. The authors attempted to prove cell
viability using LIVE/DEAD staining and 2', 7'-bis-(2-carboxyethyl)-5-(and-6)-
carboxyfluorescein (BCECF-AM) staining. The LIVE/DEAD stain was ineffective because
the hydrogel fluoresced strongly green, so live cells could not be observed above the
background fluorescence. Additionally, the BCECF-AM stain indicated enzyme activity, not
true viability, because enzymes remain active after a cell is no longer considered viable. This
work did not develop a field-deployable sensor with the photopolymer, but simply tried to
prove viability within the matrix.
24
One difficulty associated with the photopolymerizable systems is mixing the cells
with the gel precursor solution because the components are highly viscous and the shear
stresses on the cells during mixing may increase cell mortality. Modifications to the gel
structure have been undertaken in an attempt to create a low molecular weight hydrogel
based on alginate. Kong et al. (2003) created different molecular weight alginates by
irradiation and the viscosity of each was measured by shear stress and shear rate. Results
showed that higher radiation doses reduced the molecular weight most significantly, but
doses in the mid-range of those tested made the gels stiffer because only the flexible
connectors of the gel matrix broke during radiation, leaving the strong bonds intact. At low
doses, no change in stiffness was observed. Osteoblasts were mixed with the gels before
polymerization. The lower molecular weight gels resulted in greatly improved viability as
measured visually by microscopy using the Trypan blue stain, which cannot enter cells with
intact membranes, but stains the nuclei of cells with compromised membranes.
Overall, the photopolymerizable system is preferable for microfluidic work because it
can be formed into a variety of shapes by changing the photomask prior to polymerization.
Photopolymerizable polymers have been used to successfully immobilize both eukaryotic
and prokaryotic cells, as well as to immobilize enzymes for the creation of a sensor.
However, the major challenges that users of photopolymerizable polymers face are the
toxicity of the polymer components and the difficulty visualizing cells within the polymer
microscopically, because the polymer tends to absorb stains such as the green stain of the
LIVE/DEAD package.
25
Scope of Research
Based on this review of biosensors developed previously, it is clear that the need for a
non-engineered bacterial sensor for a range of toxic compounds has not yet been addressed.
Many groups have created compound-specific sensors using both whole cells as well as
purified enzymes, but no general stress response sensor has been developed yet. Microtox®
uses a non-engineered strain, but quantifies toxicity, not stress. In wastewater treatment,
electrophilic stress results in process upset and reduced treatment capacity. To reduce
instances of process upset, a sensor that responds to electrophilic insult quantified by
monitoring the intrinsic stress response of the immobilized cells, not by engineering a known
response into the cells, can be developed.
The objective of this work is to evaluate the performance of an immobilized bacterial
culture that will comprise the biological sensing element within a biosensor for detecting
electrophilic compounds based on activation of the GGKE mechanism. The biosensor will
consist of bacterial cells immobilized in a hydrogel matrix in a microfluidic flow through
setup. In its ultimate application, the biosensor will direct water to be monitored over the
immobilized cells. Any electrophilic toxins in the flow will trigger potassium efflux from the
cells. The potassium concentration will be measured using an ion-selective film (Kopelman
et al., 1997) over which the flow will pass both before and after contacting the cells. The
change in potassium will be monitored and, if efflux is indicated, the device will implicate
the presence of electrophilic toxins. For wastewater treatment applications, electrophilic
toxins have been linked to deflocculation and, thus, to process upset. A positive response by
the sensor would give plant operators early warning of process upset potential in the influent
so that the toxic slug could be rerouted to storage and slowly fed in for treatment to avoid
26
upset. For surface, ground, or drinking water applications, many chemical agents that could
be used to threaten homeland security are classified as electrophiles, such as nerve agents,
and could be effectively sensed. On the whole, the sensor under development would improve
process stability at wastewater plants and improve confidence in water supply at drinking
water plants.
To determine the feasibility of a biosensor based on the intrinsic potassium efflux
stress response, the potassium efflux potential of polymer-immobilized bacterial culture was
evaluated experimentally by comparing to non-immobilized planktonic cultures.
Furthermore, a model was developed to determine of the number of cells needed to efflux
detectable levels of potassium, while estimating the oxygen requirement for these cells to
determine if oxygen would be limiting within the system. Three matrices, alginate, a
photopolymerizable polymer, and a thermally reversible gel, were evaluated as possible
hydrogels for immobilization. Both the viability and the potassium efflux from immobilized
cells were elucidated. This work will become the basis for the incorporation of the
immobilized cells into the microfluidic device.
27
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34
Evaluating strategies for integrating bacterial cells into a biosensor designed to detect process upset
Part A: viability and activity
K. Linares*, D. Fleming**, Y. Xu***, N. Love*, B. Love**, K. Meehan*** Departments of *Civil & Environmental Engineering, **Materials Science and Engineering,
and *** Electrical and Computer Engineering, Virginia Tech, Blacksburg, VA 24061
Planned for submission to Sensors and Actuators B Abstract: The biological element of a biosensor to predict process upset in wastewater treatment plants is characterized. The bacterial culture was immobilized in a range of hydrogel polymers: a calcium alginate, a series of photopolymerizable polymers, and a thermally-reversible gel. Oxygen uptake rates and LIVE/DEAD staining were used to determine the activity and viability of the immobilized cultures. The alginate showed good viability, but poor mechanical stability in that the matrix dissolved after five days. The photopolymerizable polymers resulted in high mortality, although the matrix remained stable. The thermal polymer showed extended viability up to 12 days after immobilization, and the material did not deteriorate with time. 1. Introduction
Biosensors show promise for real time monitoring of toxicity in environmental
samples. A biosensor is generally defined as an analytical tool that pairs a biological element
with a detection system to produce a measurable response, which can be quantified into the
amount of analyte present in a sample (Belkin, 2003). The response is typically measured by
electrochemical or optical means. Biological elements are typically enzymes, antibodies,
proteins, nucleic acids, or whole cells (Belkin, 2003). Whole cells have a certain limited
lifespan, have longer response times than isolated components, and may show significant
variability between cell batches (Ramanathan et al., 2000). On the other hand, whole cells
are less expensive to use because their components require no isolation and purification as in
the cell component assay systems. Also, whole cells provide information on the
bioavailability of toxins being monitored instead of simply reporting the concentration
present. Overall, whole cell biological elements provide a means to monitor more elaborate
biological responses than biosensors based on in vitro biochemical responses.
35
For general environmental monitoring, many compounds may be responsible for the
toxicity of a sample, and so development of sensors that indicate many different toxic
compounds are required. Creating a general response biosensor is typically approached by
developing a panel of bacteria, each responsive to a different threat. Panels of E. coli
bacteria were engineered to create sensors for heat shock, oxidative stress, and protein
damage (Belkin et al., 1997), DNA damage (Belkin, 2003), and oxidative damage,
membrane damage, DNA damage, and protein damage (Kim and Gu, 2003). A field-ready
biological element was developed by Premkumar et al. (2003), who immobilized bacteria
that were engineered to report on heat shock, oxidative stress, fatty acids, peroxides, and
genotoxicity in thick silicate films, which maintained viability and activity for over a month
(Premkumar et al., 2003).
On the whole, genetically engineered strains used in biosensors have not been
commercialized to a large degree. The difficulty in overcoming the genetically modified
organism (GMO) regulations makes such sensors primarily useful only in laboratory settings,
where the threat of accidental release is minimized. Despite their advantages in terms of
robust, repeatable responses, engineered strains have yet to be widely commercialized.
Non-engineered cells do not mandate such strict controls on release as those that have
genetic modifications. While engineered cells may yield a more robust response, wild-type
cells facilitate eventual commercialization of a sensor, exemplified by the Microtox® assay,
which uses naturally luminescent Vibrio fischerii bacteria. Additionally, the use of
indigenous, wild-type cells may more accurately reflect the conditions under observation
than the use of an engineered lab strain. Using wild-type cells means the intrinsic cell
36
processes must be harnessed to detect the compounds of interest or determine if the cells are
inhibited.
The purpose of this work was to explore strategies for immobilizing non-engineered
bacterial cells for inclusion into a biosensor designed to predict process upset in wastewater
treatment plants. Process upset is commonly characterized by such symptoms as ineffective
biological oxygen demand (BOD) removal, ineffective nitrification, deflocculation, and
sludge bulking (Love and Bott, 2000). These process effects observed during an upset event
result in poor treatment and possibly permit violations. Developing a biosensor based on the
natural cell physiological response to chemical insult would give an early warning that toxic
chemicals were present. This warning would significantly benefit wastewater treatment
operations by allowing operators time to respond to the approaching threat, and could also
clarify the type of operational adjustment needed to prevent or remediate treatment process
damage.
For this study, bacterial cells were immobilized in a range of polymer matrices and
cell viability and activity was assessed. Immobilization of the bacterial cells was important
because it keeps the cell concentration relatively constant by maintaining cell viability while
reducing cell division. Additionally, immobilized cells cannot slough off as a biofilm layer
and clog downstream channels in the device. Such clogging is particularly problematic if a
liquid microfluidic conveyance system is used. Chemically polymerized hydrogels, such as
sodium alginate, are useful for macroscale work and form beads from the gel precursor
solution (Drury et al., 2004). Photopolymerized polymers have also been used for
microfluidic applications because they can easily be formed into a variety of shapes in situ by
using various photomasks to control the areas of the precursor that are polymerized (Beebe et
37
al., 2000; Heo et al., 2003). Further, viable cells have been immobilized within
photopolymers. One type of photopolymer, polyethylene glycol (PEG) diacrylate (PEG-DA)
with Darocure 1173 as the photoinitiator polymerized with UV light, has been used to
immobilize E. coli cells (Heo et al., 2003) within a microfluidic channel constructed of
polydimethylsiloxane (PDMS). The authors attempted to prove cell viability using
LIVE/DEAD staining, but found the stain ineffective because the hydrogel fluoresced
strongly green, so live cells could not be observed above the background fluorescence.
Overall, photopolymers have been used for cell immobilization because it is easy to form
them into a variety of shapes in situ, while the alginate is suited only for macroscale
applications because the ability to control the shape of the polymerized product is limited.
In the current work, a set of three polymers was investigated for use as the
immobilization matrix in the biological element of the biosensor: a calcium alginate, a series
of photopolymerizable polymers, and a thermally-reversible gel. The specific goals of the
work were to evaluate viability and activity of immobilized cells to determine a preferred
matrix for further study of detectable physiological responses to chemical perturbations.
2. Materials and Methods
2.1 Bacterial culturing
2.1.1 Growth conditions
Pseudomonas aeruginosa was isolated previously from a local wastewater treatment
plant and was grown in a mineral salt medium denoted PA M9. This medium consisted of
NaH2PO4, 3.0 g; Na2HPO4*7H2O, 6.0 g; NH4Cl, 1.0 g; NaCl, 0.50 g; MgSO42-*7H2O, 0.246
g; CaCl2, 0.0147g; FeSO4*7H2O, 2.5 mg; ZnCl2, 0.25 mg; MnSO4*H2O, 0.185 mg; CuSO4,
0.030 mg; NaMoO4*2H2O, 0.006 mg; CoCl2*6H2O, 0.001mg; H3BO3, 0.03 mg; glacial
38
acetic acid, 0.89 mL per liter solution. A second medium denoted PA BT was used to
improve alginate stability when it was the immobilizing hydrogel. The PA BT media
consisted of: NaH2PO4, 0.3 g; Na2HPO4*7H2O, 0.6 g; NH4Cl, 0.1 g; NaCl, 0.05 g; Bis Tris,
1.0 g; MgSO42-*7H2O, 0.246 g; CaCl2, 0.0147g; FeSO4*7H2O, 2.5 mg; ZnCl2, 0.25 mg;
MnSO4*H2O, 0.185 mg; CuSO4, 0.030 mg; NaMoO4*2H2O, 0.006 mg; CoCl2*6H2O,
0.001mg; H3BO3, 0.03 mg; and glacial acetic acid, 0.89 mL per liter solution. After
preparing media but before autoclaving, the pH was adjusted to 7.0 using 50% w/v NaOH
(approximately 3 mL per liter PA M9 media, and about 4.5 mL per liter PA BT media).
Escherichia coli strain K-12, ATCC 29947, was grown on mineral salt medium M9.
This media consisted of NaH2PO4, 3.0 g; Na2HPO4*7H2O, 6.0 g; NH4Cl, 1.0 g; NaCl, 0.50 g;
MgSO42-*7H2O, 0.246 g; CaCl2, 0.0147g; Thiamine HCl, 1.0 mg; and D-glucose, 2.0 g per
liter solution. No pH adjustment was required for this media. All media components were
obtained from Fisher Scientific (Pittsburgh, PA).
Potassium stock was eliminated from the culture media to reduce the dilution required
for the potassium efflux experiments. Potassium contamination from the sodium phosphate
buffers, sodium chloride, and sodium hydroxide appeared to be adequate for growth
requirements, and resulted in media concentration of about 2 mg/L.
Cultures were maintained in 250 mL Erlenmeyer flasks using a gyratory water bath
shaker (Model G76, New Brunswick Scientific, Edison, NJ) set at 37oC and were transferred
to fresh media every two days using a 1:100 dilution factor. To expand the volume of
culture, cells were transferred at the appropriate dilution to a five liter jug that was aerated
using an aquarium pump and diffuser with magnetic stirring placed in an incubator at 37oC.
Monthly, cultures were restarted from frozen stock by streaking frozen culture on Luria-
39
Bertani (LB) agar plates consisting of: NaCl, 10g; tryptone/peptone, 10 g; yeast extract, 5 g;
and Bacto agar, 10 g per liter solution. The streaked plate was incubated overnight at 37oC.
Then, a single colony was placed aseptically into fresh sterile media.
2.1.2 Growth Curve Preparation
A growth curve was determined for each strain using sidearm flasks containing 100
mL growth media incubated in a rotary water bath shaker maintained at 37oC. Transmittance
through the media was measured at 590 nm on a spectrophotometer (Spectronic 20, Bausch
and Lomb, Philadelphia, PA). Separate growth curves were determined
spectrophotometrically for cultures grown in five liter batches.
2.2 Immobilization polymer preparations
2.2.1 Alginate immobilization
The calcium alginate hydrogel matrix, a copolymer of β-D-mannuronate (M-residue)
and α-L-guluronate (G-residue) (reviewed in Rehm 1998), was used as the preliminary
immobilization matrix for the cells because this matrix has been used extensively by others
(for example, Elasri et al., 2000; Kohler, 2000; Polyak et al., 1997, 2001; Webb et al., 1997).
To form alginate beads, five liters P. aeruginosa were grown overnight to mid-log growth
state. The cells were concentrated by centrifugation in multiple bottles for 20 minutes at
4420 x g and the supernatant was discarded. The cells were resuspended in about 20 mL of
fresh media and divided into aliquots of equal cell number for the immobilization. The
resuspended bacterial concentrate was added to a 2% solution (w/v) of sodium alginate
(Protanal LF10/60, FMC BioPolymer, Philadelphia, PA, USA) in water using approximately
a 10:1 (v/v) ratio of alginate to cell solution. The solution was well mixed and slowly
40
dropped by a 21-gauge syringe into a 10% solution (w/v) of calcium chloride in water to
form spheres. The beads were rinsed with nanopure water and placed in PA BT media.
PA BT media was used for alginate bead experiments because the beads were found
to be unstable in the PA M9 media. The central modification for the BT media was the
substantial reduction in sodium concentration. Because the alginate polymerizes by
substitution of calcium for sodium within the polymer matrix, it was thought that ion
exchange could compromise alginate stability and return the beads to the liquid state, which
occurred within an hour in the PA M9 media.
The number of cells per batch was determined by dividing the cell concentration by
the number of beads per batch. Cell concentration was determined by plating triplicate serial
dilutions in sterile nanopure water of the resuspended culture on Luria-Bertani (LB) agar.
The plates were incubated overnight at 37oC, and the cell count was determined by counting
the number of colonies per plate and correcting for the dilution factor.
2.2.2 Photo-polymer immobilization
The photopolymerization procedure made it very difficult to create immobilizing
beads; therefore, flat disks approximately 0.5 cm in diameter and 0.1 cm in thickness were
formed instead. The resuspended bacterial concentrate was added to one of two versions of
the photopolymer:
• Photo 1 contained a pre-mixed solution of 5% (w/v) bis(2,4,6-
trimethylbenzoyl)-phenylphophineoxide (Irgacure 819, Ciba, Suffolk, VA,
USA) with poly(ethylene glycol) (PEG) (MW 400) containing acrylate ends
(SR 344, Sartomer, Exton, PA, USA) mixed with the bacterial concentrate to
yield a 25% (w/v) polymer solution.
41
• Photo 2 contained a pre-mixed solution of 1% (w/v) Irgacure 819 with PEG
mixed with the bacterial concentrate to yield a 22 % (w/v) polymer solution.
For both compositions, the cells were concentrated into a volume of about 8 mL of
media as described in Section 2.2.1, and the concentrated cell solution was mixed with the
polymer precursor. The mixture was dropped by pipette onto a plastic sheet and cured for 10
minutes using an aquarium lamp with peak emission at 420 nm. The cured disks were
removed from the sheet and rinsed with nanopure water, and then placed in PA M9 media.
A third photopolymer composition, Photo 3, was created using a different initiator.
This solution contained the initiator 2-hydroxy-1-[4-(2-hydroxyethoxy)phenyl]-2-methyl-1-
propanone (sold as Irgacure 2959, Ciba, Suffolk, VA, USA) in the Irgacure 819 PEG
monomer base. The solutions were premixed as 1% (w/v) of Irgacure 2959 in PEG. The
bacterial concentrate was then added and mixed as described above to yield a 22% (w/v)
polymer solution. Curing was performed using the same lamp with a cure time of 15
minutes. The disks were rinsed and placed in PA M9 media immediately afterwards. The
cell concentration per batch was calculated as described in Section 2.2.1.
2.2.3 Thermal polymer immobilization
A thermal polymer was also tested. As with the photopolymer, small beads were
difficult to generate. To form the thermal polymer sphere, the cells were concentrated into
about 20 mL of media as described in Section 2.2.1. The concentrated cell solution was
mixed at room temperature in a scintillation vial with a 15% (w/v) solution of liquid thermal
polymer, N-isopropylacrylamide-co-acrylic acid (NIPA-co-Aac), a copolymer with 98 mole
percent NIPA and 2 mole percent AAc, to result in a 10% (w/v) polymer solution. The vial
was equilibrated in a water bath at 40oC for about five minutes until the mixture solidified
42
into a white gel-like sphere about 1.5 cm in diameter. The somewhat spherical gel was
removed from the vial and wrapped in single mesh fabric (21 holes per inch) to facilitate
handling. The gel was then placed in the gyratory water bath shaker in PA M9 media at 40oC
to maintain the polymer in the gelled state. The cell concentration in the matrix was
determined by cell counts, as described in Section 2.2.1.
2.3 Cell activity and viability experiments
Bacterial activity and viability within the planktonic and immobilized cultures were
evaluated respirometrically using oxygen uptake rates and microscopically using a
LIVE/DEAD® stain, respectively. The LIVE/DEAD® stain has been widely used to
determine the viability of cells in a population by comparing the number that stain red,
representing dead cells, to the number that stain green, representing live cells.
2.3.1 Oxygen uptake rates
Oxygen uptake rates (OURs) were used to determine the activity of planktonic and
immobilized bacteria over time, with the procedure shown schematically in Figure 1.
Measurements were performed with planktonic cultures in parallel with immobilized cultures
at time zero, and continued with immobilized cultures over time. For alginate-immobilized
cultures, beads were prepared and wrapped in mesh fabric (21 holes per inch) to facilitate
handling. Beads were maintained aerobically under sterile conditions in 250 mL Erlenmeyer
flasks containing 100 mL of PA BT media, and were transferred to fresh media daily. To
determine the OUR, the mesh-bundled beads were transferred from the growth flask to a 300
mL BOD bottle, which was brought to a volume of 300 mL with media and sealed with the
dissolved oxygen (DO) probe (Orion 97-08-99, Beverly, MA). The decrease in dissolved
oxygen concentration was monitored over time using a computerized data acquisition system
43
(LabView version 6.0, National Instruments, Austin, TX) with data collected by an attached
meter (Accument Research AR25 Dual Channel ph/Ion Meter, Fisher Scientific, Pittsburgh,
PA). After the oxygen uptake rate analysis, the mesh-bundled beads were transferred to a
sterile flask with fresh media so that measurements could be performed at a later time using
the same immobilized cells. All SOUR tests were performed in duplicate.
To determine OURs for the photopolymer immobilized cells, the disks were placed in
300 mL BOD bottles and PA M9 media was added. The bottles were sealed with the DO
probes, and the experiment proceeded as described above.
To determine OURs for the thermal polymer immobilized cells, the fabric-bundled
gel was removed from its maintenance flask and transferred to a 300 mL BOD bottle. PA
M9 media at 40oC was added to the bottles, which were sealed with the DO probes. The test
proceeded as above, except that the bottle was immersed in a 40oC water bath for the
duration of the SOUR test. To compensate for the temperature variation, a planktonic control
tested at day zero was evaluated at the same temperature.
2.3.2 LIVE/DEAD® staining
The LIVE/DEAD® BacLight™ Bacterial Viability Kit (Molecular Probes, Eugene,
Oregon, USA) was used with a fluorescent microscope (Zeiss Axioscope 2 Plus, Thornwood,
NY, USA) and camera system (Zeiss AxioCam MRm, Thornwood, NY, USA) to visually
compare the numbers of viable and nonviable cells at 20 and 40x magnification. The kit
contents were mixed 1:1 and three microliters of the mixture were added to each milliliter of
cells to be stained. To stain all gels, an aliquot of the gel was placed in a microcentrifuge
tube with 1 mL of water and stained with the same amount of dye.
44
3. Results and Discussion
3.1 Oxygen uptake rates (OURs) show extended activity in thermal gels, low activity in
photopolymer, and activity until material failure in alginate gels
OURs performed on cells immobilized in the first formulation of the photopolymer
(Photo 1) showed near-zero activity after polymerization, as shown in Figure 2. Therefore,
the concentration of initiator was reduced as much as possible without compromising curing
quality or time. This new formulation, which contained 1% rather than 5% initiator, was
polymer Photo 2. Unfortunately, this reduction still resulted in very low activity (Figure 2).
Therefore, the initiator was changed to Irgacure 2959 at 1% (w/v) (polymer Photo 3), but the
results continued to show low activity (Figure 2). Changing the cell type may improve the
activity, but the current combination of cells and photo-matrices resulted in an immediate and
severe reduction in cell respirometric activity when immobilized in the matrix.
In comparison, cell activity was good in the alginate and thermal matrices, although
the OUR results indicated more limited activity initially as compared to planktonic controls
containing the same number of cells. Figure 2 summarizes the OUR results from these
matrices relative to planktonic controls measured at t=0. Because several groups show
elevated OURs relative to control, oxygen diffusion into the polymers appeared uninhibited.
In the alginate, the increased OURs probably indicate cell division and colonization within
the structure, as observed by Mater et al. by microscopy of alginate beads containing
immobilized Pseudomonas strains. This colonization occurs within the pores of the alginate
matrix, which are large enough to permit cell division. The pore size of the thermal polymer
is unknown. Because the OURs in the thermal gel also increase over the control, cell
division may also be occurring within the polymer structure. In the alginate matrix, the rates
45
increase consistently, which supports the hypothesis that the cells are using the alginate as a
carbon source, creating space for cell division as they consume the structure within which
they are contained. Perhaps because of this consumption, the matrix deteriorated within a
week of bead manufacture, preventing quantification of the long-term viability of the
alginate-immobilized cells. However, an experiment performed to determine whether the
cells could use alginate as a sole carbon source showed no difference in initial oxygen uptake
rates between immobilized cells (0.050 mg DO per cell per minute) and planktonic cells
(0.045 mg DO per liter per minute), both in media without the acetate carbon source. By day
two, the OUR for immobilized cells had dropped to 0.019 mg DO per cell per minute, much
lower than control, indicating that the cells were losing activity because the carbon source
was unacceptable. Perhaps the alginate is metabolized along with HAC, but alone the cells
do not consume the polymer.
The thermal polymer showed excellent material stability, with no deterioration
observed for over two weeks. The oxygen uptake rates fluctuate relative to the control, as
shown in Figure 2, but on the whole the cells remained viable. Clearly, the thermal polymer
is a good choice for the immobilization matrix because it is nontoxic to the cells and supports
extended viability. However, integrating the thermal polymer into the biosensor is a
challenge because of the temperature requirements of this matrix. To overcome this
constraint, the sensor requires a small heater to maintain the temperature at the appropriate
level (for example, by incorporating titanium/platinum films to permit temperature
measurement and regulation, as reviewed in Erickson and Li, 2004). Alternately, the
polymer could be modified to shift the critical temperature to a range below ambient (about
25οC). In either case, the thermal polymer showed the most promise for cell activity and,
46
thus, for inclusion in the biosensor.
3.2 LIVE/DEAD® staining is effective in determining immobilized viability in thermal
polymer
The stain was applied to cells immobilized in all matrices tested, but was only
effective for the thermal polymer. The spherical bead structure of the alginate prevented the
microscope camera from focusing on a single plane of bacteria. Attempts to flatten to beads
to permit adequate observation showed a strong background fluorescence with the green
stain, making observations of live cells difficult (Figure 3). The photopolymer matrix also
fluoresces very strongly green, making observations of live cells difficult, as shown in Figure
3. The thermal polymer showed the ideal response to the dye, in that zero background
fluorescence was observed after allowing the polymer to liquefy at room temperature.
The staining of the thermal and photopolymers reinforces the results obtained from
the OUR experiments. As shown in Figure 3, all the bacteria immobilized in the
photopolymer stain red, whether or not the cells were dropped on the surface of the polymer
or mixed with it prior to polymerization. The images shown are typical of those from the
other photopolymer formulations. Because the background fluorescence is so strongly green,
one could argue that live cells are present that cannot be observed with this method. While
that is possible, the extremely low OURs combined with the number of red cells observed in
the photopolymers support the conclusion that the survivability in the matrix was very low.
The thermal polymer shows good viability, even after 10 days within the matrix, because
green cells dominate over red. On the whole, these results replicate the SOUR data,
showing that the photopolymer has low viability, while the thermal polymer cells remain
mostly viable.
47
4. Conclusions
A series of experiments were conducted in this work to identify the most appropriate
polymer matrix for bacterial cell immobilization for a biosensor. The ideal matrix is
physically and chemically stable over several weeks, non-toxic to the cells, and will not limit
the diffusion of oxygen, compounds necessary for cell growth, or compounds, which would
trigger a stress response. Three matrices were examined here: a calcium alginate, several
photopolymerizable hydrogels, and a thermal polymer.
The sodium alginate matrix was unstable, although bacterial consumption of the
matrix as a substrate was not validated as the cause. The material degraded within a week,
rendering it incapable of containing the bacteria. The photopolymer was toxic to the cells,
resulting in significant cell death immediately after polymerization. Changes to the
concentration and the composition of the initiator did not reduce the matrix toxicity. Finally,
the thermal polymer matrix was non-toxic, stable, and did not hinder oxygen diffusion. The
only limitation on the incorporation of the thermal polymer within the microfluidic sensor is
the need to maintain a higher-than-ambient temperature in the cell chamber (40oC).
However, adding a micro-scale heater such as that reviewed in Erickson and Li (2004) would
meet this need. Alternatively, the polymer could be modified to change its immobilization
temperature. By modifying the polymer structure, the critical temperature could be reduced
below ambient, and thus reduce the need for a heating element. On the whole, even without
such chemical modifications the thermal polymer shows great potential for incorporation in
the biosensor.
5. Acknowledgements
48
The project was supported by the Paul L. Busch Award and the EPA Midwest
Hazardous Substances Research Center. KAL acknowledges the support of the Charles E.
Via, Jr. Department of Civil and Environmental Engineering Endowment. The authors
would like to thank Julie Petruska, Jody Smiley, and Phil Wunderly for analytical and
laboratory support.
49
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Beebe, David J., Jeffrey S. Moore, Joseph M. Bauer, Qing Yu, Robin H. Liu, Chelladural Devadoss, and Byung-Ho Jo. (2000) Functional hydrogel structures for autonomous flow control inside microfluidic channels. Letters to Nature 404 588-590 Belkin, Shimshon. (2003) Microbial whole-cell sensing systems of environmental pollutants. Current Opinion in Microbiology 6 206–212 Belkin, Shimshon, Dana R. Smulski, Sara Dadon, Amy C. Vollmer, Tina K. Van Dyk, and Robert A. Larossa. A panel of stress-responsive luminous bacteria for the detection of selected classes of toxicants. (1997) Water Research (31) 12, 3009-3016 Drury, Jeanie L., Robert G. Dennis, David J. Mooney. (2004) The tensile properties of alginate hydrogels. Biomaterials 24, 3187-3199 Elasri, Mohamed O., Tricia Reid, Steven Hutchens, Robert V. Miller. (2000) Response of a Pseudomonas aeruginosa biofilm community to DNA-damaging chemical agents. FEMS Microbiology Ecology 33, 21-25 Erickson, David, and Dongqing Li. (2004) Integrated microfluidic devices. Analytica Chimica Acta 507, 11–26 Gu, Man Bock, and Sue Hyung Choi. (2002) A portable toxicity biosensor using freeze-dried recombinant bioluminescent bacteria. Biosensor and Bioelectronics. 17, 433-440 Heim S, Schnieder I, Binz D, Vogel A, Bilitewski U. (1999) Development of an automated microbial sensor system. Biosens Bioelectron 14, 187–93 Heo, Jinseok, K. Joseph Thomas, Gi Hun Seong, and Richard M. Crooks. (2003) A microfluidic bioreactor based on hydrogel-entrapped E. coli: cell viability, lysis, and intracellular reactions. Anal. Chem. 75(1) 22-26 Horsburgh, Alison M., D.P. Mardlin, N.L. Turner, R. Henkler, N. Strachan, L.A. Glover, G.I. Paton, K. Kilham. (2002) On-line microbial biosensing and fingerprinting of water pollutants. Biosensors and bioelectronics 17, 495-501 Kim, Byoung Chan, and Man Bock Gu. (2003) A bioluminescent sensor for high throughput toxicity classification. Biosensors and Bioelectyronics 18, 1015-1021 Kohler, Sabine, Till T. Bachmann, Jutta Schmitt, Shimshon Belkin, Rolf D. Schmid. (2000) Detection of 4-chlorobenzoate using immobilized recombinant Escherichia coli reporter strains. Sensors and Actuators B 70, 139–144 Liu, Jing and Bo Mattiasson. (2002) Microbial BOD sensors for wastewater analysis. Water Research 36, 3786–3802
50
Love, N. G., Bott, C.B. (2000). WERF Project 99-WWF-2 Report: A review of and needs survey of upset early warning devices. Water Environment Research Foundation. Mater, Denis D.G., Jose E. Nava Saucedo, Nicole Truffaut, Jean-Noel Barbotin, Daniel Thomas. (1999) Conjugative plasmid transfer between Pseudomonas strains wtihin alginate bead microcosms: effect of the internal gel structure. Biotechnology and Bioengineering 65(1), 34-42 Philp, Jim C., Severine Balmand, Eva Hajto, Mark J. Bailey, Siouxsi Wiles, Andrew S. Whiteley, Andrew K. Lilley, Janos Hajto, and Sandra A. Dunbar. (2003). Whole cell immobilized biosensor for toxicity assessment of a wastewater treatment plant treating phenolics-containing waste. Analytica Chimica Acta 487(1) 61-74 Polyak, B., E. Bassis, A. Novodvorets, S. Belkin, and R.S. Marks. (1997) Optical fiber bioluminescent whole-cell microbial biosensors to genotoxicants. Water Science and Technology. 42 (12), 305-311 Polyak, Boris S., Efim Bassis, Alex Novodvorets, Shimshon Belkin, Robert S. Marks. Bioluminescent whole cell optical fiber sensor to genotoxicants: system optimization. (2001) Sensors and Actuators B 74, 18-26 Premkumar, J. Rajan, Rachel Rosen, Shimshon Belkin, and Ovadia Lev. (2002) Sol–gel luminescence biosensors: Encapsulation of recombinant E. coli reporters in thick silicate films. Analytica Chimica Acta 462, 11–23 Ramanathan, S., M. Ensor, and S. Daunert. (1997) Bacterial biosensors for monitoring toxic metals. Trends Biotechnol. 15, 500-506 Rehm, Bernd H.A. (1998) Alginate lyase from Pseudomonas aeruginosa CF1/M1 prefers the hexameric oligomannuronate as substrate. FEMS Microbiology Letters 165, 175-180 Reid, Brian J., Kirk T. Semple, Christopher J. Macleod, Jedda J. Weitz, Graeme I. Paton. (1998) Feasibility of using prokaryote biosensors to assess acute toxicity of polycyclic aromatic hydrocarbons. FEMS Microbiology Letters 169, 227-233 Tan, T.C., and C. Wu. (1999) BOD sensors using multi-species living or thermally killed cells of a BODSEED microbial culture. Sens Acutators B 54, 252–260 Taurianinen, Sisko, Matti Karp, Wei Chang, and Marko Virta. (1998) Luminescent bacterial sensor for cadmium and lead. Biosensor and Bioelectronics 13, 931-938 Webb O. F., P.R. Beinkowski, U. Matrubutham, F.A. Evans, A. Heitzer, G.S. Sayler. (1997) Kineticis and response of a Pseudomonas fluorescens HK44 biosensor. Biotechnology and Bioengineering. 54 (5), 491-502
51
Figure 1: Diagram of the OUR experiment protocol. Cells were grown up in 5 L jugs, concentrated by centrifugation, resuspended, and an aliquot was taken for plate counts. Next, the concentrated cells were divided into four equal batches, with two sets being immobilized in the polymer, and two remaining in the planktonic state. Here, alginate immobilization is shown as an example. The oxygen uptake rates were determined using oxygen probes and a computerized data acquisition system in duplicate for the immobilized cells as well as the planktonic cells.
52
Bead Age (days)
-2 0 2 4 6 8 10 12
Oxy
gen
Upt
ake
Rat
e Pr
opor
tiona
l to
Con
trol
0.0
0.2
0.4
0.6
0.8
1.0
1.2
1.4
1.6
1.8
Photo 1 Photo 2 Photo 3 Alginate Thermal
Rate less than control
Rate greater than control
Figure 2: OUR tests for cells immobilized in each polymer matrix. Data is normalized
to a planktonic control with equal cell density by dividing the observed rate within the gels by the average rate observed in the controls at t=0. Photopolymers show near zero oxygen uptake, while thermal and alginate matrices show extended viability. The alginate deteriorated within a week, so further testing with that matrix was impossible.
53
A B
C D
Figure 3: Representative LIVE/DEAD® stains of immobilized cells in photopolymer Photo 1 (A and B), alginate (C), and thermal polymer (D). Cells were dropped on the surface of the polymer before polymerization in Image B, while in A cells were completely mixed with the polymer before polymerization. Photopolymer and alginate images were taken immediately after polymerization, while Image D shows the thermal polymer 10 days after immobilization.
54
Evaluating strategies for integrating bacterial cells into a biosensor designed to detect electrophilic toxins
Part B: potassium efflux
K. Linares*, D. Fleming**, Y. Xu***, N. Love*, B. Love**, K. Meehan*** Departments of *Civil & Environmental Engineering, **Materials Science and Engineering,
and *** Electrical and Computer Engineering, Virginia Tech, Blacksburg, VA 24061
Planned for submission to Sensors and Actuators B Abstract: The biological element of a biosensor to detect electrophilic compounds by harnessing the glutathione-gated potassium efflux (GGKE) stress response system through the detection of effluxed potassium is characterized. The potassium efflux caused by a model electrophile is elucidated in planktonic cultures of Psuedomonas aeruginosa and Escherichia coli. P. aeruginosa was selected for further study because it is an environmentally relevant strain and effluxed comparable potassium per cell to E. coli. The bacterial culture was immobilized in two hydrogel polymers: a calcium alginate and a thermally-reversible gel. The potassium efflux from immobilized cells caused by N-ethyl maleimide, a model electrophile, was elucidated. The efflux from the alginate and thermal polymers was strong initially, but deteriorated with time, probably due to potassium limitation in the media. 1. Introduction
Real-time monitoring of toxicity in environmental samples is primarily performed
through biosensors or bioassays, because laboratory analytical techniques take more time and
require knowledge of the compound of interest in the sample. A number of whole-cell
biosensors have been developed using genetically modified bacterial strains that respond to
particular compounds, such as phenol (Philp et al., 2003; Gu and Choi, 2002), polycyclic
aromatic hydrocarbons (PAHs) (Reid et al., 1998), and heavy metals (Taurianinen et al.,
1998). One drawback with most whole cell biosensors is that they employ genetically
modified bacteria as the reporter, the use of which is prohibited in many locations. One
notable exception is the commercial Microtox® assay, which uses naturally luminescent
Vibrio fischerii bacteria. Non-engineered cells do not mandate strict controls on release
whereas those that have genetic modifications are heavily regulated. Additionally, the use of
indigenous wild type cells more accurately reflects the conditions under observation than do
55
engineered lab strains. With wild-type cells the intrinsic cellular physiology must be
harnessed to detect the presence of compounds of interest or to determine if the cells are
inhibited or stressed.
Electrophiles are a class of compounds that have been shown to trigger measurable
stress responses in bacterial cells (Bott and Love, 2002). Industrially relevant electrophilic
(oxidative) toxins include a range of chemicals, such as heavy metals and organic compounds
containing chloro- and imide- constituents. This class of toxins, commonly found in
industrial wastewater, is believed to impair biomass function in biological treatment plants
when present at shock loads, and can result in process upset (Bott and Love, 2002). The
most common modes of process upset are ineffective biological oxygen demand (BOD)
removal, ineffective nitrification, deflocculation, and sludge bulking (Love and Bott, 2000).
Although there is substantial speculation about the origin of these upsets, operators generally
have insufficient information to identify the source of the problem. However, when asked,
operators suggested that heavy metals and toxic organic compounds (which include
electrophilic chemicals) are among the most common sources of these upsets (Love and Bott,
2000).
It is speculated that electrophilic chemicals cause process upset by triggering stress
response mechanisms in the biomass. Activated sludge deflocculation, one common
component of process upset, has been linked to bacterial activation of a stress response called
the glutathione-gated potassium efflux (GGKE) response (Bott and Love, 2002). This
mechanism is initiated when an electrophile enters the cell and the reaction between the
electrophile and glutathione (N-(N-L-γ-glutamy-L-cysteinyl)glycine) activates potassium
efflux pumps located in the cell membrane (Apontoweil and Berends, 1975 a and b) (shown
56
in Figure 1). GGKE transports potassium ions out of the cell in exchange for hydrogen ions
transported into the cell, which acidify the cytoplasm and cause activation of secondary stress
responses that protect cells from oxidative damage (Ferguson et al., 2000).
In biomass, the GGKE response results in an increase in extracellular potassium,
adding additional monovalent cations in the floc matrix, and increasing the intrafloc
monovalent to divalent cation ratio (Bott and Love, 2002). This ratio is an important
predictor of floc strength (Higgins and Novak, 1997 a and b). Disrupting that ratio makes
flocs unstable and results in loss of biomass from the system (Murthy et al., 1998). This type
of upset encompasses the modes of upsets observed with electrophilic shock, namely,
ineffective BOD removal, deflocculation, and sludge bulking. The development of sensors
around this GGKE response to detect electrophilic compounds entering sewers or wastewater
treatment plants would improve process operation and effluent quality by predicting the
process effect linked to such upset events, and thus allow the adjustment of process
operations to accommodate and minimize the process impact.
The goal of this project was to develop a biosensor based on the GGKE stress
response system. By monitoring bulk potassium as an indicator of the stress experienced by
the cells, the biosensor will monitor for electrophilic compounds that have been shown to
cause deflocculation in the activated sludge process. The microscale biosensor under
development includes a whole cell biological element, potassium detection membranes, and
microfluidic componentry that compose a lab-on-a-chip device.
Within the biological element, bacterial cells will be immobilized in a polymer
matrix. Immobilization of the bacterial cells is important because it as compared to a
biofilm, it keeps the cell concentration more constant by maintaining the viability of the
57
immobilized cells while reducing cell division. Additionally, immobilized cells cannot
slough off as a biofilm layer could and clog the microfluidic channels in the device.
The focus of this work was on the design and characterization of the biological
element of the biosensor, with the specific goal of quantifying the impact of immobilization
on the rate and degree of potassium efflux in response to a model electrophile. In the current
work, two polymers were investigated for use as the immobilization matrix: a calcium
alginate and a thermally-reversible gel. In the completed sensor, the concentration of
potassium will be monitored as an indicator of bacterial stress. By utilizing the intrinsic
response of wild type cells and monitoring the change in potassium ion efflux from them, the
device is devoid of genetically modified cells and its eventual use and commercialization are
simplified greatly.
2. Materials and Methods
2.1 Bacterial culturing
2.1.1 Growth conditions
Pseudomonas aeruginosa was isolated previously from a local wastewater treatment
plant and was grown in a mineral salt medium denoted PA M9. This medium consisted of
NaH2PO4, 3.0 g; Na2HPO4*7H2O, 6.0 g; NH4Cl, 1.0 g; NaCl, 0.50 g; MgSO42-*7H2O, 0.246
g; CaCl2, 0.0147g; FeSO4*7H2O, 2.5 mg; ZnCl2, 0.25 mg; MnSO4*H2O, 0.185 mg; CuSO4,
0.030 mg; NaMoO4*2H2O, 0.006 mg; CoCl2*6H2O, 0.001mg; H3BO3, 0.03 mg; glacial
acetic acid, 0.89 mL per liter solution. A second medium denoted PA BT was used to
improve alginate stability when it was the immobilizing hydrogel. The PA BT media
consisted of: NaH2PO4, 0.3 g; Na2HPO4*7H2O, 0.6 g; NH4Cl, 0.1 g; NaCl, 0.05 g; Bis Tris,
1.0 g; MgSO42-*7H2O, 0.246 g; CaCl2, 0.0147g; FeSO4*7H2O, 2.5 mg; ZnCl2, 0.25 mg;
58
MnSO4*H2O, 0.185 mg; CuSO4, 0.030 mg; NaMoO4*2H2O, 0.006 mg; CoCl2*6H2O,
0.001mg; H3BO3, 0.03 mg; and glacial acetic acid, 0.89 mL per liter solution. After
preparing media but before autoclaving, the pH was adjusted to 7.0 using 50% w/v NaOH
(approximately 3 mL per liter PA M9 media, and about 4.5 mL per liter PA BT media).
Escherichia coli strain K-12, ATCC 29947, was grown on mineral salt medium M9.
This media consisted of NaH2PO4, 3.0 g; Na2HPO4*7H2O, 6.0 g; NH4Cl, 1.0 g; NaCl, 0.50 g;
MgSO42-*7H2O, 0.246 g; CaCl2, 0.0147g; Thiamine HCl, 1.0 mg; and D-glucose, 2.0 g per
liter solution. No pH adjustment was required for this media. All media components were
obtained from Fisher Scientific (Pittsburgh, PA).
Potassium stock was eliminated from the culture media to reduce the dilution required
for the potassium efflux experiments. Potassium contamination from the sodium phosphate
buffers, sodium chloride, and sodium hydroxide appeared to be adequate for growth
requirements, and resulted in media concentration of about 2 mg/L.
Cultures were maintained in 250 mL Erlenmeyer flasks using a gyratory water bath
shaker (Model G76, New Brunswick Scientific, Edison, NJ) set at 37oC and were transferred
to fresh media every two days using a 1:100 dilution factor. To expand the volume of
culture, cells were transferred at the appropriate dilution to a five liter jug that was aerated
using an aquarium pump and diffuser with magnetic stirring placed in an incubator at 37oC.
Monthly, cultures were restarted from frozen stock by streaking frozen culture on Luria-
Bertani (LB) agar plates consisting of: NaCl, 10g; tryptone/peptone, 10 g; yeast extract, 5 g;
and Bacto agar, 10 g per liter solution. The streaked plate was incubated overnight at 37oC.
Then, a single colony was placed aseptically into fresh sterile media.
59
2.1.2 Growth Curve Preparation
A growth curve was determined for each strain using sidearm flasks containing 100
mL growth media incubated in a rotary water bath shaker maintained at 37oC. Transmittance
through the media was measured at 590 nm on a spectrophotometer (Spectronic 20, Bausch
and Lomb, Philadelphia, PA). Separate growth curves were determined
spectrophotometrically for cultures grown in five liter batches.
2.2 Immobilization polymer preparations
2.2.1 Alginate immobilization
The calcium alginate hydrogel matrix, a copolymer of β-D-mannuronate (M-residue)
and α-L-guluronate (G-residue) (reviewed in Rehm, 1998), was used as the preliminary
immobilization matrix for the cells because this matrix has been used extensively by others
(for example, Elasri et al., 2000; Kohler, 2000; Polyak et al., 1997, 2001; Webb et al., 1997).
To form alginate beads, five liters P. aeruginosa were grown overnight to mid-log growth
state. The cells were concentrated by centrifugation in multiple bottles for 20 minutes at
4420 x g and the supernatant was discarded. The cells were resuspended in about 20 mL of
fresh media and divided into aliquots of equal cell number for the immobilization. The
resuspended bacterial concentrate was added to a 2% solution (w/v) of sodium alginate
(Protanal LF10/60, FMC BioPolymer, Philadelphia, PA, USA) in water using approximately
a 10:1 (v/v) ratio of alginate to cell solution. The solution was well mixed and slowly
dropped by a 21-gauge syringe into a 10% solution (w/v) of calcium chloride in water to
form spheres. The beads were rinsed with nanopure water and placed in PA BT media.
PA BT media was used for alginate bead experiments because the beads were found
to be unstable in the PA M9 media. The central modification for the BT media was the
60
substantial reduction in sodium concentration. Because the alginate polymerizes by
substitution of calcium for sodium within the polymer matrix, it was thought that ion
exchange could compromise alginate stability and return the beads to a liquid state, which
occurred within an hour in the PA M9 media.
The number of cells per batch was determined by dividing the cell concentration by
the number of beads per batch. Cell concentration was determined by plating triplicate serial
dilutions in sterile nanopure water of the resuspended culture on Luria-Bertani (LB) agar.
The plates were incubated overnight at 37oC, and the cell count was determined by counting
the number of colonies per plate and correcting for the dilution factor.
2.2.2 Thermal polymer immobilization
A thermal polymer was also tested. As with the photopolymer, small beads were
difficult to generate. To form the thermal polymer immobilization gel, the cells were
concentrated into about 20 mL of media as described in Section 2.2.1. The concentrated cell
solution was mixed at room temperature in a scintillation vial with a 15% (w/v) solution of
liquid thermal polymer, N-isopropylacrylamide-co-acrylic acid (NIPA-co-Aac), a copolymer
with 98 mole percent NIPA and 2 mole percent AAc, to result in a 10% (w/v) polymer
solution. The vial was equilibrated in a water bath at 40oC for about five minutes until the
mixture solidified into a white gel-like sphere about 1.5 cm in diameter. The somewhat
spherical gel was removed from the vial and wrapped in single mesh fabric (21 holes per
inch) to facilitate handling. The gel was then placed in the gyratory water bath shaker in PA
M9 media at 40oC to maintain the polymer in the gelled state. The cell concentration in the
matrix was determined by cell counts, as described in Section 2.2.1.
61
2.3 Potassium Efflux Experiments
Potassium efflux experiments at various time points on the growth curve with
planktonic cultures were carried out by growing five liters P. aeruginosa or E. coli overnight
to the desired growth state in the appropriate growth medium. The cells were concentrated
by centrifugation in multiple bottles for 20 minutes at 4420 x g and the supernatant was
discarded. The cells were resuspended in fresh media, combined, and the volume was
adjusted to 700 mL with fresh media. Samples were taken from the well mixed flask for
soluble potassium and plate counts. The resuspended culture was divided evenly among six
flasks with 100 mL per flask. The flasks were placed on a multiple position stir plate and
aerated using an aquarium pump to ensure oxygenation during the experiment. One milliliter
automatic pipetter tips were placed on the end of the air tubing in each flask to reduce bubble
size and to prevent contamination of the tubing. Each of three flasks was shocked with 1 mL
of a 5 mg/mL N-ethyl malemide (NEM) stock, resulting in a dose of 50 mg/L of NEM.
Three control flasks remained undosed with NEM. Samples for potassium measurement
were taken over a one-hour time period by filtering the media through 0.2 µm nitrocellulose
MCE filters (25 mm diameter, Fisher Scientific, Pittsburgh, PA). Samples were acidified
with concentrated nitric acid for preservation, and then diluted 1:10 with nanopure water.
One milliliter of diluted sample was removed and replaced with 1 mL of 12.7 g/L cesium
chloride stock solution (Alfa Aesar, Ward Hill, MA) to minimize interference from sodium.
Potassium standards were prepared using a potassium reference solution (Fisher Scientific,
Fairlawn, NJ) and the same concentration of cesium chloride was added to standards. The
samples were analyzed on an Atomic Absorption Spectrometer (AA) (5100 PC Atomic
Absorption Spectrometer, Perkin Elmer, Norwalk, CT). All glassware used in the potassium
62
measurements was prepared by acid washing in 10% nitric acid, followed by triple rinsing
with nanopure water. Error for potassium efflux potential was calculated from the standard
deviation of averages for analysis from three flasks and included standard deviation from the
cell counts. Cell counts were determined by plating serial dilutions in sterile nanopure water
of the resuspended culture on Luria-Bertani (LB) agar. The plates were incubated overnight
at 37oC, and the cell count was determined by counting the number of colonies per plate and
correcting for the dilution factor.
Potassium efflux experiments with immobilized cultures (procedure shown in Figure
2) were carried out by growing 5 L culture overnight to mid-log growth state (absorbance of
the media was between 0.10 and 0.20 at 590 nm) in PA BT media. The polymer
immobilization was carried out as described in Section 2.2. Once complete, the immobilized
cells were equally divided among six flasks containing 100 mL media each. The experiment
on the potassium concentration effluxed per cell after a shock with NEM proceeded in the
same manner as described for the planktonic samples, but the experiment time was extended
to two hours.
2.4 Alginate bead material stability testing
The material stability of the alginate beads after exposure to equivalent
concentrations of potassium to that observed during efflux was determined using a texture
analyzer (TA-XT2i Texture Analyzer, Texture Technologies Corporation, Scarsdale, NY).
The instrument was operated to measure force in compression, with a test speed of 0.1mm/s
using the 5 kg load cell. Rupture was deemed as the peak of the force curve before reaching
the stage.
63
To determine the effect of potassium on the bead stability, alginate beads without
immobilized cells were incubated for two weeks in deionized water containing potassium at a
range bracketing the values that would be observed per bead from cell efflux. The maximum
efflux per cell was scaled down based on the cell loading per bead to determine the
potassium dose per bead. Ten beads were used for each concentration, and the
concentrations selected were 0.3 and 0.6 mg potassium per 1.1 mL of liquid to bracket the
possible range of efflux potential. The control consisted of 10 beads in 1.1 mL deionized
water with no added potassium.
3. Results and Discussion
3.1 Planktonic efflux experiments show no difference between bacterial strains
A series of potassium efflux experiments were conducted on planktonic cultures of P.
aeruginosa and E. coli across their growth curves to determine which strain effluxed the
most potassium per cell, and to determine which growth state elicits the strongest efflux
response. A typical result from a batch efflux experiment is shown in Figure 3. Figure 4
shows the series of batch experiments that were performed to compare the potassium efflux
potential of the two strains at various growth states on media containing comparable
potassium concentration. The average K+ efflux in planktonic cultures grown to mid log
state was (4.50 + 0.4) x 10-10 mg K+ per cell for P. aeruginosa and (5.54 + 0.8) x 10-10 mg K+
per cell for E. coli, while cells in late log phase averaged a potassium efflux of (3.07+ 0.4) x
10-10 mg K+ per cell for P. aeruginosa and (4.15 + 0.3) x 10-10 mg K+ per cell for E. coli.
Although it appears that late log phase (0.20-0.25 and 0.25-0.30 absorbance units measured
at 590 nm for E. coli and P. aeruginosa, respectively) experiments produced somewhat lower
efflux potential than mid log phase cells (0.025-0.20 and 0.025-0.25 absorbance units
64
measured at 590 nm for E. coli and P. aeruginosa, respectively) no difference exists in the
average efflux potentials based on a statistical t-test between mid and late log phase
experiments with either culture. Further, no difference exists between the average efflux
potentials between the cultures at either growth state. Since the activated sludge P.
aeruginosa isolate used in these experiments effluxes comparable K+ on average per cell, we
continued to use P. aeruginosa in further experiments because it may more accurately
represent conditions at wastewater plants than the laboratory E. coli strain.
3.2 Material failure in alginate beads cannot be attributed to K+ efflux
Because the alginate matrix degraded after only five days of polymerization,
experiments were conducted to determine if the potassium efflux from the cells caused the
breakdown of the polymer structure. The alginate polymerizes by substitution of calcium for
sodium within the matrix, and matrix stability was hypothesized to be affected by re-
equilibration in sodium. Because of the similarity in the properties of sodium and potassium,
it was thought that the potassium effluxed from the cells interacts with the matrix and aids in
its degradation. However, the concentration of potassium that would be effluxed into the
bulk liquid had no effect on the bead stability as measured by a texture analyzer to measure
force in compression, as shown in Figure 5.
3.2 Potassium efflux from alginate-immobilized cells is reduced as compared to
planktonic cultures, but nearly equal for thermal polymer-immobilized cells
Experiments to determine the amount of potassium effluxed per cell while
immobilized were carried out with P. aeruginosa at mid-log growth state in calcium alginate
beads. Sample results from initial efflux experiments conducted with alginate-immobilized
cells are shown in Figure 6. A t-test showed that the potassium concentration at point four
65
was statistically different from the initial value, indicating that efflux occurred. The efflux
potential with the immobilized culture was (1.3 + 0.1) x 10-10 mg K+ per cell, as shown in
Table 1, which is much lower than the efflux from planktonic cultures. However, efflux was
significantly reduced in experiments conducted on the same immobilized cells after five days
(shown in Figure 7), probably because the maintenance media had a very low potassium
concentration, and the cells were unable to replenish their internal potassium stocks. The
large variability and the reduced initial efflux potential observed in the data from the alginate
immobilized cells is likely due to electrophilic diffusion limitations into the beads and K+ out
of the beads, although these limitations were not explored or quantified.
On the other hand, the efflux potential from the thermal polymer-immobilized cells
was higher than that from the alginate and comparable to the results for planktonic culture, as
shown in Table 1. The efflux observed one day after immobilization was (4.6 + 0.8) x 10-10
mg K+/cell, four times greater than that of the alginate-immobilized cells (Figure 8). As in
the alginate, the efflux potential decreased with time. Figure 9 shows the efflux results from
cells immobilized for five days. The figure shows no clear efflux trend, although the control
seems to be uptaking potassium. Again, it is thought that the reduced efflux potential with
time is attributed to the low-potassium maintenance media.
Table 1: The efflux potential from immobilized P. aeruginosa in two polymer matrices on day one as compared to the average of four planktonic experiments
Matrix Efflux Potential (mg K+ / cell) n
None (Planktonic) (4.50 + 0.4) x 10-10 4
Alginate (1.3 + 0.1) x 10-10 1
Thermal (4.6 + 0.8) x 10-10 1
66
4. Conclusions
A series of experiments were conducted in this work to select an appropriate polymer
matrix for bacterial cell immobilization within the biosensor under development. The ideal
matrix will be physically and chemically stable over several weeks, will be non-toxic to the
cells, and will not limit the diffusion of oxygen, compounds necessary for cell growth, or the
electrophiles which would trigger a stress response. Two matrices were examined here: a
calcium alginate and a thermal polymer.
Both polymers showed stable initial efflux results, but later efflux experiments on the
polymers resulted in unstable and reduced efflux, perhaps because the potassium
concentration in the matrix was limited, thus preventing the cells from replenishing their
potassium stocks. The sodium alginate matrix was materially unstable and unable to contain
the bacterial cells after five days, but the cause could not be attributed to the potassium
effluxed from the cells. Efflux from the alginate beads was four times lower than that from
the thermal or the planktonic cells, and the variability was much higher. The thermal
polymer was non-toxic, stable, and the initial efflux potential from thermal polymer-
immobilized cells was nearly equal to that from planktonic cells. The only limitation on the
incorporation of the thermal polymer within the microfluidic sensor is the need to maintain a
higher-than-ambient temperature in the cell chamber. On the whole, the thermal polymer
shows great potential for incorporation in the biosensor.
5. Acknowledgements
The project was supported by the Paul L. Busch Award and the EPA Midwest
Hazardous Substances Research Center. KAL acknowledges the support of the Charles E.
Via, Jr. Department of Civil and Environmental Engineering Endowment. The authors
67
would like to thank Julie Petruska, Jody Smiley, and Phil Wunderly for analytical and
laboratory support.
68
References
Apontoweil, P., Berends, W. (1975a). Glutathione biosythesis in Escherichia coli K-12: properties of the enzymes and regulation. Biochimica et Biophysica Acta, 399 (1), 1-9. Apontoweil, P., Berends, W. (1975b). Isolation and initial characterization of glutathione-deficient mutants of Escherichia coli K-12. Biochimica et Biophysica Acta. 399(1), 10-22. Bott, C. B. and Love, N. G. (2002) Investigating a mechanistic cause for activated sludge deflocculation in response to shock loads of toxic electrophilic chemicals. Water Environment Research. 74, 306-315. Bott, Charles B. (2001) Elucidating the role of toxin-induced microbial stress responses in biological wastewater treament process upset. Diss. Virginia Polytechnic Insitute and State University. Elasri, Mohamed O., Tricia Reid, Steven Hutchens, Robert V. Miller. (2000) Response of a Pseudomonas aeruginosa biofilm community to DNA-damaging chemical agents. FEMS Microbiology Ecology 33, 21-25 Erickson, David, and Dongqing Li. (2004) Integrated microfluidic devices. Analytica Chimica Acta 507, 11–26 Ferguson, G., McLaggan P. D., Booth, I.R. (1995). Potassium channel activation by glutathione-S-conjugates in Escherichia coli. Molecular Microbiology. 17(6), 1025-1033. Gu, Man Bock, and Sue Hyung Choi. (2002) A portable toxicity biosensor using freeze-dried recombinant bioluminescent bacteria. Biosensor and Bioelectronics. 17, 433-440 Higgins, M.J. and Novak, J.T. (1997a) Dewatering and settling of activated sludges: The case for using cation analysis. Water Environment Research 69 (2), 225-232
Higgins, M.J. and Novak, J.T. (1997b) The effect of cations on the settling and dewatering of activated sludges: Laboratory results. Water Environment Research 69 (2), 215-224 Love, N. G., Bott, C.B. (2000). WERF Project 99-WWF-2 Report: A review of and needs survey of upset early warning devices. Water Environment Research Foundation. Kohler, Sabine, Till T. Bachmann, Jutta Schmitt, Shimshon Belkin, Rolf D. Schmid. (2000) Detection of 4-chlorobenzoate using immobilized recombinant Escherichia coli reporter strains. Sensors and Actuators B 70, 139–144 Murthy, SN, JT Novak, RD De Haas. (1998) Monitoring cations to predict and improve activated sludge settling and dewatering properties of industrial wastewaters. Water Science and Technology. 38 (3), 119-126
69
Philp, Jim C., Severine Balmand, Eva Hajto, Mark J. Bailey, Siouxsi Wiles, Andrew S. Whiteley, Andrew K. Lilley, Janos Hajto, and Sandra A. Dunbar. (2003). Whole cell immobilized biosensor for toxicity assessment of a wastewater treatment plant treating phenolics-containing waste. Analytica Chimica Acta 487(1) 61-74 Polyak, B., E. Bassis, A. Novodvorets, S. Belkin, and R.S. Marks. (1997) Optical fiber bioluminescent whole-cell microbial biosensors to genotoxicants. Water Science and Technology. 42 (12), 305-311 Polyak, Boris S., Efim Bassis, Alex Novodvorets, Shimshon Belkin, Robert S. Marks. Bioluminescent whole cell optical fiber sensor to genotoxicants: system optimization. (2001) Sensors and Actuators B 74, 18-26 Rehm, Bernd H.A. (1998) Alginate lyase from Pseudomonas aeruginosa CF1/M1 prefers the hexameric oligomannuronate as substrate. FEMS Microbiology Letters 165, 175-180 Reid, Brian J., Kirk T. Semple, Christopher J. Macleod, Jedda J. Weitz, Graeme I. Paton. (1998) Feasibility of using prokaryote biosensors to assess acute toxicity of polycyclic aromatic hydrocarbons. FEMS Microbiology Letters 169, 227-233 Taurianinen, Sisko, Matti Karp, Wei Chang, and Marko Virta. (1998) Luminescent bacterial sensor for cadmium and lead. Biosensor and Bioelectronics 13, 931-938 Webb O. F., P.R. Beinkowski, U. Matrubutham, F.A. Evans, A. Heitzer, G.S. Sayler. (1997) Kineticis and response of a Pseudomonas fluorescens HK44 biosensor. Biotechnology and Bioengineering. 54 (5), 491-502
70
Figure 1: The glutathione-gated potassium efflux (GGKE) system. When an electrophile enters a cell, it can damage DNA and proteins by oxidation. Cells have developed the GGKE stress response to reduce this type of damage. Reduced glutathione, GSH, reacts with the electrophile to create a glutathione-electrophile conjugate. The presence of this conjugate within the cells triggers efflux channels that efflux potassium outside the cell while concurrently importing hydrogen ions. These hydrogen ions acidify the cytoplasm, and the pH drop affords protection from the electrophile for the DNA and proteins within the cell. Overall, the GGKE stress response system is characterized by potassium efflux in response to an electrophilic challenge. (After Dr. Charles Bott, 2001)
71
Figure 2: The experimental setup for potassium efflux experiments. Culture was grown to the appropriate growth state in a 5 L jug, then concentrated by centrifugation and resuspended. An aliquot for plate counts was removed. The concentrated culture was divided among six flasks, which were aerated and, at time=0, dosed with NEM. Samples for potassium were taken over time to determine the increase in potassium concentration due to cell efflux.
72
Time (min)
0 10 20 30 40 50 60 70
Pota
ssiu
m C
once
ntra
tion
(mg/
L)
0
1
2
3
4ControlDosed
Figure 3. Batch K+ efflux experiment with planktonic P. aeruginosa at an absorbance (measured at 590 nm) of 0.215 after exposure to 50 mg/L N-ethylmaleimide (NEM) versus an unshocked control. Error bars represent the standard deviation of the averages for three flasks.
73
Absorbance
0.00 0.05 0.10 0.15 0.20 0.25 0.30 0.35
Pota
ssiu
m E
fflux
per
cel
l (m
g K+ /c
ell)
0
2e-10
4e-10
6e-10
8e-10
1e-9P. aeruginosaE. coli
Mid log Late log
Figure 4. K+ efflux potentials (mg K+ effluxed per cell) versus planktonic culture
density, measured as absorbance (at 590 nm), for P. aeruginosa and E. coli.
74
Figure 5: No significant difference is observed in the force required to rupture for
alginate beads incubated over time with potassium concentrations comparable to that effluxed from immobilized cells.
75
Time (min)
0 20 40 60 80 100 120 140
Pota
ssiu
m C
once
ntra
tion
(mg/
L)
0
1
2
3
4ControlDosed
Figure 6. K+ efflux ((1.3 + 0.1) x 10-10 mg K+ per cell) at t=1 day from P. aeruginosa immobilized in calcium alginate beads after exposure to 50 mg/L NEM versus an unstressed, immobilized control.
76
Time (min)
0 20 40 60 80 100 120 140
Pota
ssiu
m C
once
ntra
tion
(mg/
L)
-1
0
1
2
3
4
5
ControlDosed
Figure 7: K+ efflux ((1.3 + 0.1) x 10-10 mg K+ per cell) at t=5 days from P. aeruginosa
immobilized in calcium alginate beads after exposure to 50 mg/L NEM versus an unstressed, immobilized control.
77
Time (min)
-20 0 20 40 60 80 100 120 140
Pota
ssiu
m C
once
ntra
tion
(mg/
L)
0
2
4
6
8
10
ControlDosed
Figure 8. K+ efflux ((4.6 + 0.8) x 10-10 mg K+/cell ) at t=1day from P. aeruginosa immobilized in thermal polymer gels after exposure to 50 mg/L NEM versus an unstressed, immobilized control.
78
Time (min)
-20 0 20 40 60 80 100 120 140
Pota
ssiu
m C
once
ntra
tion
(mg/
L)
0
2
4
6
8
10
12
14
ControlDosed
Figure 9: K+ efflux ((4.6 + 0.8) x 10-10 mg K+/cell ) at t=5 days from P. aeruginosa
immobilized in thermal polymer gels after exposure to 50 mg/L NEM versus an unstressed, immobilized control.
79
Chapter 4: Engineering Significance
Engineering Significance
Properly functioning wastewater treatment plants are vital to human health and water
quality. However, even the most efficient plant may have instances of process upset,
characterized by deflocculation and poor nutrient and BOD removal, that result in the failure
of the treatment train. During such an event, operators typically are unaware that upset has
begun until the event is well underway and the damage to the biological treatment step has
been done. Providing operators with advanced warning when an upset is about to occur
would allow them to initiate protective measures to ensure the continued functioning of the
treatment process. Such advance warning would be possible with the biosensor under
development here.
This biosensor will be the first to harness the GGKE stress response mechanism to
link the efflux of potassium from immobilized cells with the presence of an electrophile in
the sample. The change in potassium concentration across the sensor caused by potassium
efflux from immobilized cells will be monitored; and, at a certain threshold increase,
activation of the GGKE system will be inferred. Because the system is activated by
electrophiles, the sensor will not be useful for upset events caused by other classes of
compounds, such as oil and grease. Since many upsets can be linked to electrophilic shock
and potassium efflux and, thus, to deflocculation, the sensor will serve a vital role in
improving the functionality of wastewater treatment plants under the influence of toxic
chemical shocks.
Incorporating this sensor in the real-time analyses of plant influent will inform
operators of potential upset due to electrophilic shock. However, it will not tell the whole
80
story of the potential for upset events because it is only useful for predicting electrophilic
shock upset. Combining this sensor with other real-time sensors, such as sensors measuring
the influent’s effect on oxygen uptake rates within the bioreactor, would provide a more
complete picture of the upset potential of the influent. Adding sensors for other effects on
the biomass to create a sensor network that reponds to plant influent in real time would
significantly improve the operator’s control over process operations by providing information
on the types of effects the influent may cause in the biomass. Although such a sensor would
provide limited information on the cause of the effect, the early warning allows operators
time to reroute the suspect influent or initiate treatment steps to mitigate the toxic effects of
the influent.
In addition to using an array of biosensors to monitor the plant influent, such sensors
might be applied to test the effluent from industries that discharge to the sewer system. This
testing could be conducted as a prerequisite for discharge to the sewer, to ensure the safety of
the large quantities of wastewater generated from industrial activities. Alternately, daily
samples of industrial wastewater could be collected and the testing could be performed on the
stored samples after an upset event, or after the upset warning sensors at the plant were
triggered, in an attempt to find the cause of the upset. Such a system would work well in an
area containing a few high volume industrial contributions where the testing of stored
samples would take minimal time. As the number of industries increases, the time to
determine the source of the cause increases, as does the age of the sample tested, reducing the
reliability of the results. Clearly, for such attempts to link the causative industry to an upset
event, the results of the biosensor must be proven and reliable. Extensive tests to determine
the response of the sensor against background shifts in pH and temperature must be
81
performed to rule out the possibility of false positive results. On the whole, an array of
biosensors to monitor for process upset by electrophilic shock and by other modes of upset
would improve the state of wastewater treatment by allowing operators time to respond to the
threat, and by potentially enabling them to link the each upset with its cause.
82
Appendix A: Data for Chapter 2 A.1 Data for oxygen uptake rate experiments Data averaged from Probe A and B Presented in Chapter 2, Figure 2 A.1.1 Photopolymerizable polymer: Photo 3, 4-21-04, Book 2, Page 87
83
Oxygen Uptake Rates for Photopolymer Gel with New Initiator (Photo 3): 4-21-04
y = -0.3153x + 7.2672R2 = 0.9912
y = -0.0066x + 7.1144R2 = 0.9595
0
1
2
3
4
5
6
7
8
0 5 10 15 20 25 30Time (min)
Dis
solv
ed O
xyge
n (m
g/L)
Immobilized
Planktonic
A.1.2 Photopolymerizable polymer: Photo 2, 3-18-04, Book 2, Page 77
84
Oxygen Uptake Rates for Photopolymer Gel, Photo 2, 3-18-04
y = -0.116x + 7.4596R2 = 0.9948
y = -0.0103x + 7.3975R2 = 0.6349
0
1
2
3
4
5
6
7
8
0 5 10 15 20 25 30Time (min)
Dis
solv
ed O
xyge
n (m
g/L)
Planktonic
Immobilized
85
A.1.3 Photopolymerizable polymer: Photo 1, 2-26-04, Book 2, Page 72
86
Oxygen Uptake Rates for Photopolymer Gel, Photo 1, 2-26-04
y = -0.3574x + 7.418R2 = 0.9965
y = -0.0064x + 7.211R2 = 0.673y = -0.0013x + 6.8977
R2 = 0.0168
0
1
2
3
4
5
6
7
8
0 5 10 15 20 25Time (min)
Dis
solv
ed O
xyge
n (m
g/L)
Immob 1Immob 2
Planktonic
A.1.4 Alginate oxygen uptake rates Alginate polymer: day zero, 4-18-04, Book 2, Page 84
87
88
Alginate polymer: days 1, 3, and 5
89
Oxygen Uptake Rates for Alginate Gels: Experiment Begun 4-18-04
y = -0.0933x + 7.9481R2 = 0.9938
y = -0.102x + 8.3643R2 = 0.9983
y = -0.1479x + 8.5671R2 = 0.9995
y = -0.0775x + 7.5802R2 = 0.9951
y = -0.149x + 8.2379R2 = 0.996
0
1
2
3
4
5
6
7
8
9
0 5 10 15 20 25 30 35Time (min)
Dis
solv
ed O
xyge
n (m
g/L)
PlanktonicInitial
ImmobInitial
ImmobDay 1
ImmobDay 3
ImmobDay 5
90
A.1.5 Thermal polymer oxygen uptake rates Thermal polymer: planktonic control, and polymer on days 0, 1, 5, 8, 10, and 14 4-18-04, Book 2, Page 84
91
92
93
Oxygen Uptake Rates for Thermal Polymer Gels: Experiment Begun 4-18-04
y = -0.0933x + 7.9481R2 = 0.9938
y = -0.0465x + 5.5066R2 = 0.9022
y = -0.1077x + 5.4475R2 = 0.9704
y = -0.0698x + 4.7016R2 = 0.9744
y = -0.1134x + 6.9157R2 = 0.9975
y = -0.1375x + 4.324R2 = 0.9916
0
1
2
3
4
5
6
7
8
9
0 5 10 15 20 25 30 35Time (min)
Dis
solv
ed O
xyge
n (m
g/L)
PlanktonicInitial
ImmobInitial
ImmobDay 1
ImmobDay 3
ImmobDay 5
ImmobDay 12
94
A.2 Alginate as a carbon source experiment, 5-22-04, Book 2, Page 91 Referenced in Chapter 2, Section 3.1 Planktonic control with HAC
Planktonic control without HAC
95
Alginate immobilized cells, no HAC, day zero
96
Alginate immobilized cells, no HAC, day 2
97
Summary plot from alginate degradation experiment
Oxygen Uptake Rates for Alginate Gels: Experiment Begun 4-18-04
y = -0.0192x + 9.807R2 = 0.983
y = -0.0449x + 7.2751R2 = 0.9971
y = -0.0500x + 7.3609R2 = 0.9889
y = -1.4576x + 4.8813R2 = 0.9999
0
2
4
6
8
10
12
0 5 10 15 20 25 30 35 40 45 50Time (min)
Dis
solv
ed O
xyge
n (m
g/L)
Planktonicwith HAC
ImmobInitial
ImmobDay 2
Planktonicno HAC
98
A.3 t-tests to determine statistical difference t tests for planktonic potassium efflux experiments: (units = mg K+/cell): Referenced in Chapter 3 Data:
99
Appendix B: Data for Chapter 3 B.1 Growth Curve Data B.1.1 Psuedomonas aeruginosa 7-22-03, Book 2, Page 12
P. aeruginosa Growth Curve in 5L Jug
0.000
0.050
0.100
0.150
0.200
0.250
0.300
0.350
0 10 20 30 40 50 60 7
Time (hrs)
Abs
orba
nce
0
100
B.1.2 Escherchia coli 8-14-03, Book 2, Page 16
E. coli Growth Curve in 5 L Jug
0.00
0.05
0.10
0.15
0.20
0.25
0.30
0 5 10 15 20 25 30 35
Time (hrs)
Abs
orba
nce
101
B.2 Data for potassium efflux per cell for planktonic cultures B.2.1 Planktonic efflux from P. aeruginosa Summary table of efflux data for planktonic P. aeruginosa
The following sets of data were obtained by acidifying the samples with nitric acid, diluting, and analyzing on the AA. Data shown has been corrected for dilution and for machine drift during analysis. Control flasks are indicated by “C”, and dosed flasks by “F.” Potassium concentration is reported in mg/L. Potassium efflux from transmittance = 61, 6-20-03, Book 1, Page 99
Potassium efflux from transmittance = 72, 7-18-03, Book 2, Page 9
Potassium efflux from transmittance = 60, 7-19-03, Book 2, Page 10
Potassium efflux from transmittance = 50, 7-24-04, Book 2, Page 11
102
Potassium efflux from transmittance = 55, 7-29-03, Book 2, Page 14
Potassium efflux from transmittance = 91, 7-31-03, Book 2, Page 15
Potassium efflux from transmittance = 79, 11-23-03, Book 2, Page 53
Potassium efflux from transmittance = 75, 11-23-03, Book 2, Page 54
Potassium efflux from transmittance = 66, 1-2-04, Book 2, Page 59
Potassium efflux from transmittance = 62, 1-17-04, Book 2, Page 63
103
B.2.2 Planktonic efflux from E. coli Summary table of efflux data for planktonic E. coli
The following sets of data were obtained by acidifying the samples with nitric acid, diluting, and analyzing on the AA. Data shown has been corrected for dilution and for machine drift during analysis. Control flasks are indicated by “C”, and dosed flasks by “F.” Potassium concentration is reported in mg/L. Potassium efflux from transmittance = 56, 8-15-03, Book 2, Page 17
Potassium efflux from transmittance = 56, 8-20-03, Book 2, Page 18
Potassium efflux from transmittance = 78, 8-22-03, Book 2, Page 20
Potassium efflux from transmittance = 70, 8-23-03, Book 2, Page 21
104
Potassium efflux from transmittance = 55, 9-25-03, Book 2, Page 33
Potassium efflux from transmittance = 76, 12-10-03, Book 2, Page 56
Potassium efflux from transmittance = 86, 12-12-03, Book 2, Page 57
Potassium efflux from transmittance = 89, 1-6-04, Book 2, Page 60
105
B.3 Data for efflux from alginate-immobilized P. aeruginosa The following data were obtained by acidifying the samples with nitric acid, diluting, and analyzing on the AA. Data shown has been corrected for dilution and for machine drift during analysis. Control flasks are indicated by “C”, and dosed flasks by “F.” Potassium concentration is reported in mg/L. Days after immobilization are given (e.g., day 0 = immediately after immobilization and day 1 = 24 hours after immobilization). Cell counts were performed by serial dilutions and plating. Volumes shown with cell counts indicate total volume of culture after resuspension. Total cells = average cell count x resuspension volume. This total was divided equally into six flasks to give cells per flask. Efflux 2, Day 1 2-4-04, Book 2, Page 69 Presented in Chapter 3, Table 1 and Figure 6
Efflux 3, Day 5 1-27-04, Book 2, Page 67 Presented in Chapter 3, Figure 7
Data not included in thesis: 1-26-04, Book 2, Page 66 Efflux 1, Day 0
106
Efflux 2, Day 4
1-27-04, Book 2, Page 67 Efflux 1, Day 0
Efflux 2, Day 3
2-4-04, Book 2, Page 69 Efflux 1, Day 0
Efflux 2, Day 2
Efflux 3, Day 5
107
B.4 Data for thermallly reversible gel-immobilized P. aeruginosa The following data were obtained by acidifying the samples with nitric acid, diluting, and analyzing on the AA. Data shown has been corrected for dilution and for machine drift during analysis. Control flasks are indicated by “C”, and dosed flasks by “F.” Potassium concentration is reported in mg/L. Days after immobilization are given (e.g., day 0 = immediately after immobilization and day 1 = 24 hours after immobilization). Cell counts were performed by serial dilutions and plating. Volumes shown with cell counts indicate total volume of culture after resuspension. Total cells = average cell count x resuspension volume. This total was divided equally into six flasks to give cells per flask. Efflux 1, Day One, 4-19-04, Book 2, Page 86 Presented in Chapter 3, Table 1 and Figure 8
5-24-04, Book 2, Page 92 Presented in Chapter 3, Figure 9 Efflux 1, Day 3
108
Data not included in thesis: 3-31-04, Book 2, Page 82 Efflux 1, Day 1
Efflux 2, Day 3
Efflux 3, Day 7
4-18-04, Book 2, Page 84 and 86 Efflux 2, Day 3
109
Efflux 3, Day 5
110
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